Greenhouse Gases And IPM 9789387214330, 9387214338

Citation preview

GREENHOUSE GASES AND IPM

Greenhouse Gases and IPM

NASIM AHMAD Natioanl Centre for Integrated Pest Management (NCIPM) ICAR-NCIPM, New Delhi

SHAHID AHAMAD Dy. Director (Research) Directorate of Research Sher-e-Kashmir University of Agricultural Sciences and TechnologyJammu (Chatha), J.&K

2018 Educationist Press

A Division of Write & Print Publications New Delhi-110 015

Cataloging in Publication Data—DK Courtesy: D.K. Agencies (P) Ltd. Ahmad, Nasim, author. Greenhouse gases and IPM / Nasim Ahmad, Shahid Ahamad. pages cm Includes bibliographical references and index. ISBN 9789387214033 1. Greenhouse gases.. 1972- author. II. Title.

2. Pests–Integrated control.I. Ahamad, Shahid,

LCC TD885.5.G73A36 2018 | DDC 363.73874

23

©2018 AUTHOR ISBN: 978-93-87214-03-3 Publisher’s note: Every possible effort has been made to ensure that the information contained in this book is accurate at the time of going to press, and the publisher and authors/editors cannot accept responsibility for any errors or omissions, however caused. No responsibility for loss or damage occasioned to any person acting, or refraining from action, as a result of the material in this publication can be accepted by the authors/editors, the publisher or the authors/editors. The Publisher is not associated with any product or vendor mentioned in the book. The contents of this work are intended to further general scientific research, understanding and discussion only. Readers should consult with a specialist where appropriate. Every effort has been made to trace the owners of copyright material used in this book, if any. The authors/editors and the publisher will be grateful for any omission brought to their notice for acknowledgement in the future authors/editors editions of the book. All Rights reserved under International Copyright Conventions. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise without the prior written consent of the publisher and the copyright owner. Published by : Hitesh Mittal For Write & Print Publications H-13, Bali Nagar New Delhi-110 015 Phone: 011-45635684 E-mail:[email protected] Website: www.writeandprint.com Laser Typesetting : Anshu Gupta Delhi - 110 059

Printed at in India.

About the Authors

Dr. Nasim Ahmad joined ICAR-Nation Research Centre for Integrated Pest Management (NCIPM). At ICAR-NCIPM, New Delhi he has been associated with the IPM teams working on various crops specially oilseeds and pulses. He did his Master’s Degree in Botany with specialization in Plant Pathology and Ph.D. in Botany with specialization in Mycology from DDU, Gorakhpur University, U.P. After his Master’s Degree he worked in Herbarium Cryptogamae Indiae Orientalis (HCIO) at Division of Plant Pathology, Indian Agricultural Research Institute, New Delhi for five years on taxonomic studies on powdery mildews fungi. He has several new varieties and new species of powdery mildews and other fungi and a new genus of hyphomycetes fungi to his credit. Later, he along with the team members he has validated various IPM Technologies on mustard, groundnut, pigeon pea and chickpea crops. He has also been associated with microbial bio-control laboratory at ICAR-NCIPM and engaged in imparting training under the NCIPM training programmes on mass production of microbial bio-control agents’ especially associated with fungal bio-control agents and their quality control aspects. He has published several peered reviewed research papers in various national/international journals of repute.

Dr. Shahid Ahamad presently working as Deputy Director Research in the Directorate of Research. He did M.Sc. in 1996 in Plant Pathology from CSAU, Kanpur and Ph.D. in Botany in 2003 from CSJMU, Kanpur, U.P. He started his career as a Research Associate from IARI, New Delhi in 1997. Joined as Jr. Scientist (Plant Pathology) in 2001 in Sher-E-Kashmir Univ. of Agril. Sciences & Technology of Jammu, J.&K. Selected as Sr. Scientist/ Programme Coordinator in 2008. He is also working as Nodal Officer, PME Cell. He is working as Reviewers/editorial members in different national and international journals. He conferred Fellow of Society of Plant Protection Sciences (2005) IARI, New Delhi & Fellow of Indian Phytopathological Society in 2015. He wins many awards viz. Young Achiever Awards - 2011 for his outstanding contribution in the field of Plant Pathology by SADHNA, H.P.; Distinguished Service Awards (2012) by Bioved Research Society, Allahabad U.P., Councilor (2012-15) of SPPS Division of Nematology, I.A.R.I., New Delhi; Scientist of the Year Award (2013) by Society of Biological Sciences and Rural Development, Allahabad, U.P. Recently he has been elected as President of Indian Phytopathological Society (NZ) for 2017. He has been published more than 250 publications in national and international reputed journals/magazines/Newspapers including 10 books in different aspects of agriculture/ Plant Pathology.

About the Book Insects and diseases arc a major challenge to greenhouse production. I I'M is an important tool in the management of these pests. The primary goal oflPM is to optimize pest control in an economically and ecologically sound way. IPM involves the integration of cultural, physical, biological, and chemical practices to grow crops with minimal use of pesticides. Monitoring, sampling, and record keeping are used to determine when control options are needed to keep pests below an economically damaging threshold. Pest management, not eradication, is the goal of IPM. Integrated pest management (IPM). also known as integrated pest control I IPC) is a broad-based approach that integrates practices for economic control of pests. IPM aims to suppress pest populations below the economic injury level (El L). A regular monitoring program is the basis of IPM decision making, regardless of the control strategies used. By regular monitoring, a scout is able to gather current information on the identity and location of pest problems and to evaluate treatment effectiveness. This is a reference book which attempts to provide postgraduate and professional readers already familiar with the subject with a means to acquire deeper knowledge on integrated control of pests and diseases in greenhouse crops and furthermore suggest possible roads to lake in future tasks. Another decisive stimulant for this endeavor was the realization of the growing need to incorporate integrated systems of protection from arthropod pests and diseases for the thousands of hectares of protected crops in the world. The fruit, vegetable, and ornamental plant markets and the technical and economic efficiency of crop protection require these integrated control systems. The book adopts multidisciplinary approach in addressing both basic and applied aspects of integrated pest management in greenhouse crops.

Contents Foreword Preface 1. Introduction 2. IPM Scouting and Decision-making 3. Identifying Pest and Beneficial Insects on Sticky Cards 4. Integrated Pest Management and Insect Biology 5. Fungal and Bacterial Diseases 6. Diseases of Greenhouse Ornamental Crops 7. Management of Fungus Gnats 8. Managing Aphids in the Greenhouse 9. Cultural Practices 10. Managing Caterpillar Pests in and around Greenhouses 11. Managing Two-Spotted Mites in the Greenhouse 12. Managing Whiteflies in the Greenhouse 13. Managing Waste Water from Intensive Horticulture: A Wetland System 14. Pest Management for Herb Bedding Plants Grown in the Greenhouse 15. Pest Management for Vegetable Bedding Plants 16. Plant Parasitic Nematodes 17. Thrips in Greenhouse Crops - Biology, Damage and Management 18. Monitoring the Greenhouse Environment Bibliography

Index

Plants arc often exposed to various abiotic and biotic stresses. They have developed specific mechanisms lo adapt, survive and reproduce under these stresses, Together, these stresses constitute the primary cause of crop losses worldwide, reducing average yields of major crop plants increasing cost of cultivation, reducing input use efficiency and impairing the quality of produce. Current climate Change scenarios predict an increase in mean temperatures and drought thai wilt drastically affect global agriculture in the near future, A complete undemanding on physiological Md molecular mechanisms especially signaling cascades in response to abiotic and biotic stresses in tolerant plants Kill help to manipulate susceptible crop plants and increase sustain agricultural productivity in the near future. Advanced aagricultural approaches are also required for sustainable solutions to the huge global problem of ‘hidden hunger* and it may be achieved by the biofortificalion for increased micronutrient intakes and improved imcronulxient status in the food, Management of cultural practices in conjunction with use of plant bio-regulators and chemicals will give a long way in management of various abiotic and biotic stresses. The information oh pest and disease management under protected cultivation of horticultural crops is scattered, I am told thai (here is no hook which deals with pests and diseases in protected horticultural crops (fruits, vegetables and ornamentals) in detail using physical, cultural, chemical, biological, host resistance, and integrated methods. In this regard, an attempt has been made by Dr. Shahid Abamad, Deputy Director Research, Directorate of Research, SKUAST-Jarnfflu and his team to come up with a volume entitled “Green House leases and IPM” containing IS chapters

I feel that this publication shall be of immense use lo researchers, undergraduate and post graduate students along with other Slake holders who are dealing with green ecosystem. I congratulate d»e authors for their painstaking efforts in bringing out this book.

Preface Insects and diseases are a major challenge to greenhouse production. IPM is an important tool in the management of these pests. The primary goal of IPM is to optimize pest control in an economically and ecologically sound way. IPM involves the integration of cultural, physical, biological, and chemical practices to grow crops with minimal use of pesticides. Monitoring, sampling, and record keeping are used to determine when control options are needed to keep pests below an economically damaging threshold. Pest management, not eradication, is the goal of IPM. Integrated pest management (IPM), also known as integrated pest control (IPC) is a broad-based approach that integrates practices for economic control of pests. IPM aims to suppress pest populations below the economic injury level (EIL). A regular monitoring program is the basis of IPM decision making, regardless of the control strategies used. By regular monitoring, a scout is able to gather current information on the identity and location of pest problems and to evaluate treatment effectiveness. To publish a book is an arduous task. Fortunately, I am convinced that the effort of the people who have guided, in one way or another, in this book has been worthwhile. Another decisive stimulant for this endeavour was the realization of the growing need to incorporate integrated systems of protection from arthropod pests and diseases for the thousands of hectares of protected crops in the world. The fruit, vegetable and ornamental plants and the technical and economic efficiency of crop protection require these integrated control systems. The information on pest and disease management under protected cultivation of horticultural crops is very much scattered. There is no book at present which comprehensively and exclusively deals with the above aspects. This book deals with pests (insect and mite) and diseases in protected

horticultural crops (fruits, vegetables, and ornamentals) in detail using physical, cultural, chemical, biological, host resistance, and integrated methods. This is a reference book which attempts to provide postgraduate and professional readers already familiar with the subject with a means to acquire deeper knowledge on integrated control of pests and diseases in greenhouse crops and furthermore suggest possible roads to take in future tasks. Another decisive stimulant for this endeavor was the realization of the growing need to incorporate integrated systems of protection from arthropod pests and diseases for the thousands of hectares of protected crops in the world. The fruit, vegetable, and ornamental plant markets and the technical and economic efficiency of crop protection require these integrated control systems. The book adopts multidisciplinary approach in addressing both basic and applied aspects of integrated pest management in greenhouse crops. Nasim Ahmad Shahid Ahamad

1: Introduction BASICS OF GREENHOUSE A greenhouse is a generic term referring to the use of a transparent or partially transparent material supported by a structure to enclose an area for propagating or growing plants. Specifically, where the covering material is glass, the structure may be referred to as a ‘glasshouse’. A ‘greenhouse’ or ‘polyhouse’ refers to the use of plastic films or sheeting. When the enclosing material is woven or otherwise constructed to allow sunlight, moisture and air to pass through the gaps, the structure is known as a ‘shade house’ or ‘screen house’. When looking to develop or expand a greenhouse enterprise, it is important to make sure that the structures you invest in are suitable and meet your needs. The shape and design of the structure influences: • • • • • • •

the amount of light transmitted the amount of natural ventilation the useable internal space efficient use of structural materials condensation run-off heating requirements the cost. When deciding on a greenhouse design for commercial production, key factors of the greenhouse need to be considered. It is not possible to provide a definitive priority list to suit everyone, but generally, the height of the structure is critical and will have significant bearing on managing the growing environment in a range of conditions. Ventilation is also at the top of

the list and roof ventilation is superior to side wall ventilation. Active ventilation systems can also be considered. Heating is essential for controlled environment horticulture and naturally the computer control systems are critical. Covering materials, screens (thermal and insect) and evaporative cooling systems should also be carefully assessed.

TYPES OF GREENHOUSE Shape Classification of a greenhouse is according to its basic shape. Types include Gable, Flat arch, Raised dome, Sawtooth, Skillion, Tunnel. Multi-span structures Multi-span greenhouses have a surface area smaller than a number of single span greenhouses of equivalent production capacity. This results in less heat loss and significant energy savings. Substantial economies of scale and production efficiencies are also attainable using multi-span designs. Multi-spans are typically more robust in design. As a result they tend to suffer less damage during storms and gale force winds.

OTHER TYPES OF STRUCTURES Shade houses Shade houses are structures which are covered in woven or otherwise constructed materials to allow sunlight, moisture and air to pass through the gaps. The covering material is used to provide a particular environmental modification, such as reduced light or protection from severe weather conditions. The height of the structure will vary according to the type of crop being produced and may be as high as 8 metres. Shade houses are used over outdoor hydroponic systems, particularly in warmer regions. Screen houses Screen houses are structures which are covered in insect screening material instead of plastic or glass. They provide environmental modification

and protection from severe weather conditions as well as exclusion of pests. They are often used to get some of the benefits of greenhouses in hot or tropical climates. Crop top structures A crop top is a structure with a roof but which does not have walls. The roof covering may be a greenhouse covering material such as plastic or glass, or shade cloth or insect screening. These structures provide some modification of the growing environment such as protection of the crop from rain or reduction of light levels.

CLASSIFYING GREENHOUSES

Low technology greenhouses have significant production and environmental limitations.

Greenhouses are a technology based investment. The higher the level of technology used, the greater potential for achieving tightly controlled growing conditions. This capacity to tightly control the conditions in which the crop is grown is strongly related to the health and productivity of the crop. The following three categories of greenhouse have been defined to assist people in selecting the most appropriate investment for their needs and budget. Low Technology Greenhouses A significant proportion of the industry in Australia currently uses low technology structures. These greenhouses are less than 3 metres in total height. Tunnel houses, or “igloos”, are the most common type. They do not have vertical walls. They have poor ventilation. This type of structure is relatively inexpensive and easy to erect. Little or no automation is used.

While this sort of structure provides basic advantages over field production, crop potential is still limited by the growing environment and crop management is relatively difficult. Low level greenhouses generally result in a suboptimal growing environment which restricts yields and does little to reduce the incidence of pests and diseases. Pest and disease control, as a result, is normally structured around a chemical spray program. Low technology greenhouses have significant production and environmental limitations, but they offer a cost effective entry to the industry. Medium Technology Greenhouses Medium level greenhouses are typically characterised by vertical walls more than 2m but less than 4 metres tall and a total height usually less than 5.5 metres. They may have roof or side wall ventilation or both. Medium level greenhouses are usually clad with either single or double skin plastic film or glass and use varying degrees of automation.

Medium technology greenhouses offer a compromise between cost and productivity

Medium level greenhouses offer a compromise between cost and productivity and represent a reasonable economic and environmental basis for the industry. Production in medium level greenhouses can be more efficient than field production. Hydroponic systems increase the efficiency of water use. There is greater opportunity to use non-chemical pest and disease management strategies but overall the full potential of greenhouse horticulture is difficult to attain. High Technology Greenhouses

High technology greenhouses offer superior crop performance

High level greenhouses have a wall height of at least 4 metres, with the roof peak being up to 8 metres above ground level. These structures offer superior crop and environmental performance. High technology structures will have roof ventilation and may also have side wall vents. Cladding may be plastic film (single or double), polycarbonate sheeting or glass. Environmental controls are almost always automated. These structures offer enormous opportunities for economic and environmental sustainability. Use of pesticides can be significantly reduced. High technology structures provide a generally impressive sight and, internationally, are increasingly being involved in agribusiness opportunities. Although these greenhouses are capital intensive, they offer a highly productive, environmentally sustainable opportunity for an advanced fresh produce industry. Investment decisions should, wherever possible, look to install high technology greenhouses.

ORIENTATION AND SITING A GREENHOUSE The orientation of the greenhouse is typically north-south. A lot of emphasis internationally is placed on orientation to maximise light interception in the greenhouse. This is not such an issue in Australia because our light levels are much higher. Shadows cast by gutters, trusses and equipment in the roof of the greenhouse can lead to uneven light conditions in the crop. As the sun moves from the east to the west during the day, the shadows of the greenhouse structure will also move. An east-west alignment creates structural shadows in the same part of the crop through the day which can affect crop productivity and plant health in this area. Subsequently, to minimise shading

effects, greenhouses are generally oriented north-south. In southern areas of Australia, an east-west orientation may result in slightly more light transmission, but the need for cooling and ventilation is a more important factor under Australian conditions. As you go further north, there is even less difference in light transmission which ever way a greenhouse is oriented. Again, cooling and ventilation and therefore the direction of prevailing winds should be the primary consideration in orienting a structure. Crop rows are also typically aligned north-south to minimise shading within the crop. In most areas, vents will be on the east and west. The direction of prevailing winds should be taken into consideration, with structures oriented to take advantage of cooling summer breezes. Where fans are used for forced ventilation, they should be positioned to minimise any likely impact on neighbours. When siting a greenhouse, you also need to take into consideration the shading effect of vegetative screens and windbreaks. Locating greenhouses against a tree line will result in lower yields because of reduced light levels. Greenhouse covering materials near trees will also become quite dirty, further reducing light transmission. When siting a greenhouse; • Favour a property with natural visual screening; • Consider proximity to key markets; • Prevent a direct line of sight between the development and adjoining dwellings or roadways; • Locate new developments, such as additional greenhouses, behind existing structures; • Locate structures with sufficient setbacks from roadsides and boundaries; • Use landscaping, mounding and vegetation to soften the impact of the development; • Keep existing vegetation and landforms wherever practical; • Consider transport routes and the availability of labour and services; • Avoid development in areas that are visually prominent or which are

• • • •

highly exposed, such as ridgelines; Locate structures so that they follow the contours of the land; Avoid locating structures on steep slopes (greater than 1 in 5); Check potential impacts of adjacent land uses in terms of pests, diseases and weeds; Take note of adjacent sensitive areas (e.g. wetlands, waterways, native vegetation) and site greenhouses appropriately.

VENTILATION IN GREENHOUSES Good ventilation is critical in maintaining an optimal growing environment and improves the overall efficiency of a greenhouse. It is essential for both good temperature and humidity management. Cooling is critical in the Australian environment and is most commonly achieved using passive roof ventilation. The movement of hot air up and out of the roof vents, pulls in cooler air. Ventilation is also important for air circulation and replenishing carbon dioxide. Poor air circulation reduces plant activity and can lead to problems with humidity and disease management. Air movement in the greenhouse should be between 0.2 and 0.7 metres per second. If carbon dioxide levels are not maintained, plant growth is affected. Ventilation is about air exchange. Large volumes of air need to be moved during hot conditions. A greenhouse needs to be able to achieve at least 30 air changes per hour, but ideally 60 air changes per hour – one air exchange every minute – is needed to make sure the greenhouse environment can be managed in hot sunny Australian conditions. Ventilation can be achieved passively or actively. The venting capacity of greenhouses is usually described as a percentage of floor area. For example, a greenhouse with 30% roof ventilation has 0.3m2 of open vent area for every 1m2 of floor area. A greenhouse in all but the coolest areas of Australia should have a venting capacity of at least 25%, but as much as 40% is desirable. It is better to have more venting capacity than you need.

VENTS

Passive ventilation uses openings (vents) which naturally draw air through the greenhouse. Vents are the most common ventilation method used in greenhouse production. Roof ventilation is a more effective method of air exchange than side wall ventilation. Though different designs will vary in their effectiveness, in general terms, roof vents are up to 5 times more effective than side wall vents.

The height of a structure and the height of the vents significantly impact on the capacity of a vent to remove heat from the greenhouse. The natural ‘chimney effect’ of rising hot air and falling cooler air which is the basis for passive ventilation becomes truly effective above approximately 3.5 metres. A low profile greenhouse therefore, will require forced cooling to provide similar suitable conditions to a tall structure.

FANS Active ventilation is the use of equipment to force air into or out of the structure. Fans are the key method of actively venting a greenhouse. Fans can also be fitted in greenhouses to move or circulate air within the greenhouse. Circulating fans are often used inside passively ventilated structures to assist air movement when venting is minimal.

When using fans for air exchange, the most effective approach is to pull the air through the full length of the structure to avoid hot air pockets remaining. Fans placed to extract air from higher in the greenhouse are more effective for cooling than fans which are placed lower. Active ventilation systems are limited in their capacity to quickly exchange large volumes of air. If the design specifications for your greenhouse are inadequate poor air circulation through the crop can result. Under ventilated structures often have overheating problems in the crop in the middle of the greenhouse. To ensure correct capacity and installation, select the fans in consultation with the manufacturer and an independent expert. Ventilation fans generally need to have sufficient capacity to completely replace the air in the greenhouse every minute. Fans have an on-going operational cost and noise generation may pose problems in some areas. Fan efficiencies influence running costs and should be considered when purchasing. It is important to clean and maintain fans to ensure that they are functioning properly.

HEIGHT OF GREENHOUSE STRUCTURE Height is one of the most important aspects of a greenhouse. The height of a structure directly impacts on natural ventilation, the stability of the internal environment and crop management. Greenhouse structures with wall heights of at least 4 metres should be constructed wherever feasible. These structures should be built in preference to designs of lower height. The natural ‘chimney effect’ of rising hot air and falling cooler air which is the basis for passive ventilation becomes truly effective above approximately 3.5 metres.

A tall, roof ventilated greenhouse can achieve a more uniform, stable and ultimately superior growing environment for the crop. During hot weather, a taller structure avoids trapping heated, humid air around the plants. Many of the problems that are encountered in greenhouse crops can be directly attributed to the capacity to manage the growing environment. Better control of the growing environment directly impacts on how well other problems in the greenhouse can be managed. A significant proportion of yield loss in Australian greenhouse crops can be attributed to poor management of heat. The capacity for a grower to manage heat in summer is greatly improved with increasing greenhouse height. Effective management of pests and diseases using non-chemical management strategies is also dependent on good control of the growing environment and the value of height in the structure can not be overstated. Although some crops can be grown relatively well in lower profile greenhouses, taller structures are more versatile, are suitable for a wider range of crops and therefore a better long term investment.

COMPUTER CONTROL SYSTEMS, SENSORS AND MONITORING EQUIPMENT IN GREENHOUSES

Monitoring growing conditions is essential

Good crop management depends on having the right information to make necessary decisions. In the past, the grower has been the greenhouse sensor and control system – checking conditions and adjusting equipment settings as needed to optimise crop growth. To improve crop management, a numberof sensors and instruments can (and should) be used to gather information. A computer control system can then use this information to make regular adjustments to equipment settings

to optimise growing conditions. Monitoring growing conditions is essential. Even without automated control of the production system, it is not possible to make the right decisions about the crop without having the right information. Temperature and relative humidity (and/or vapour pressure deficit) need be monitored in every greenhouse. Light levels should be checked at least periodically to make sure covering materials are performing adequately, but ideally light levels need to be checked on a regular basis in order to know the optimal temperature regime for the crop. The electrical conductivity and pH of both the feed and drain solutions should be monitored in every hydroponic system. Temperature and relative humidity sensors should be placed level with the growing tip of the crop. Placing a thermometer near the door of a greenhouse might be convenient but it will not give you the information needed for producing an optimal crop. Medium and high technology greenhouses make use of a range of sensors which link into automated control systems. These systems can monitor temperature, relative humidity, vapour pressure deficit, light intensity, electrical conductivity (feed and drain), pH (feed and drain), carbon dioxide concentrations, wind speed and direction and even whether or not it is raining. The information is used to control heating, venting, fans, screens, nutrient dosing, irrigation, carbon dioxide supplementation and fogging or misting systems. Correct operation of the automatic controllers is essential to management of an optimal growing environment. Emergency alarms and backup generators may be used in case of problems or power failure due to the large investments made in producing a crop.

Sophisticated sensors are being developed to monitor plants directly

Closer monitoring of the greenhouse environment with sensors and advanced software can greatly improve yields and economic performance by optimising plant growth. The cost of automated equipment and computer control systems can generally be recovered within a couple of seasons through savings in labour and better crop production. Increasingly sophisticated sensors are being developed and adopted in commercial greenhouse operations to monitor the plants directly. Today’s growers have access to continuous measurement of a wide range of aspects of plant growth including stem diameters, sap flow rates, expansion of fruit and leaf temperatures. The integration of this information into production decisions is still new, but is rapidly providing better data about growing conditions and even assisting in the early detection of plant stress. Good control in the greenhouse is the ultimate aim of controlled environment horticulture. The most important benefit of control in the greenhouse is the efficiency and effectiveness of your management decisions. There are a lot of other benefits too which save money and result in a better crop. These include greater energy and labour efficiency, more efficient use of water and fertilisers and fewer pesticides. Better control also gives you a more uniform crop so it costs you less to sell.

HEATING GREENHOUSES Heating is used to provide optimal temperatures for crop growth and for management of the humidity in the greenhouse. Heating may be needed throughout the year, not just in winter. Heat should ideally be applied as low as possible in the greenhouse (with the exception of “grow pipes”) and distributing heat evenly is essential for optimal crop production. There are essentially two methods of heating - hot air and hot water. Hydronic heating refers to the use of a boiler to heat water which is then piped through the greenhouse. The pipes, located around the walls of the structure and/or between plant rows, radiate heat. The major costs are in the boiler and piping. A centralised hydronic heating system is generally a more efficient form of heating in greenhouses greater than 1000m2 and especially where there are several separate greenhouses. When hot water heating is

used, the boilers may be situated away from the greenhouse. This flexibility provides the opportunity to locate potentially noisy boilers away from farm boundaries to minimise disturbance to neighbours. Heated air can also be used to maintain temperatures in a greenhouse. Hot air may be directly generated in the greenhouse or the internal air can be warmed through heat exchange with an external heat source. Where combustion occurs at the source of the heat, such as in a gas fired heater, locating the heater outside the greenhouse and using a heat exchange pad to warm the internal air is recommended. This is because the combustion process can result in ethylene production and also water vapour. Ethylene can cause leaf drop and premature ripening of fruit. Increased moisture levels in the air may result in excessive humidity and condensation problems. Gas remains the primary source of energy for greenhouse heating in Australia. Oil, diesel and coal are also used. Natural gas is lower cost and clean burning. It does not require on-farm storage tanks and is typically low maintenance. Unfortunately, natural gas is not available in all areas. Bottled gas (LPG) is similar to natural gas but more expensive. Costs can be volatile and storage tanks are needed. Oil and diesel are more expensive than natural gas and because these fuels do not burn as cleanly, more boiler maintenance is needed. On-farm storage is also required. Coal is relatively low cost if it is locally available. It is more polluting than other fuel sources. Large storage areas are needed on farm as well as loading equipment. As coal does not burn as cleanly, significant boiler maintenance is needed. Greenhouse heating requirements Three ways heat is lost from the greenhouse 1. Conduction – direct movement through structural materials 2. Leakage – hot air escaping through gaps and doorways 3. Radiation – radiate energy moving directly through covering materials The main basis for heating in a greenhouse is the replacement of lost heat. Heat can be lost through conduction, leakage and radiation. Most heat is lost through conduction, where heat energy is transferred directly through

covering materials and the structure to the outside atmosphere. Leakage of air accounts for the next greatest amount of lost heat. In a well constructed and maintained structure as much as 10% of heat loss can still be due to leakage. In greenhouses with poorly fitting doors, partially opened vents, other gaps or broken covering materials, significantly more heat can be lost this way. The heat lost through radiation depends on the covering material. It is often ignored. Glass will not allow much heat to be lost through radiation, though polyethylene film is practically transparent to thermal radiation. Working out how much heating is needed Heat losses are worked out for the coldest expected night temperatures. This then gives the maximum heating capacity needed. Heater capacity is calculated from the heat load (Q) of the greenhouse and the heater efficiency. Calculating the total heat load (QT) The total heat load is the sum of the amount of heat loss through all three different processes – conduction (QC) + Leakage (QL) + Radiation (QR); QT = QC + QL + QR Radiation is the third way in which heat can be lost from the greenhouse. The heat load through radiation (QR) depends on the covering material. Glass does not permit much heat to escape through radiation, but basic polythene covering materials can. There are, however, a number of plastic films are on the market today that restrict thermal radiation. These are known as thermic films. The loss due to radiation is generally ignored, so the total heat load is; QT = QC + QL Heat load due to conduction (QC) Conduction - the transfer of heat through the structural materials – is the main way that heat is lost. Different materials have a different conduction values. These are referred to as ‘U’ values or heat transfer values. They are measured in Watts per square metre per degree Kelvin or you may find them in Btu per hour per square foot per degree Fahrenheit.

You also need to know the surface area of the greenhouse and the difference between the temperature set point for your crop and the coldest outside temperature.

Heat transfer values Covering material

U value Watts / m2 °

Btu / hr sq.ft °F

Single polythene plastic film

7.5 – 8.5

1.2 – 1.5

Double polythene plastic film

4.0 – 5.0

0.7 – 0.9

Polycarbonate

6.8

1.2

Polycarbonate double walled

3.5

0.6

Glass 3 mm

7.0 – 8.0

1.2 – 1.4

Glass 6 mm

6.5

1.15

Thermal screen

2.8

0.5

SURFACE AREA (SA) OF GREENHOUSE The amount of heat that can be transferred out of the greenhouse is dependent on the surface area of the structure. A large surface area can lose more heat than a smaller surface area over the same period of time. The surface area of the greenhouse is referred to as ‘SA’ and is measured in square metres. The area includes the roof and all the walls of the greenhouse. Gable type greenhouse For a gable type greenhouse, you need to measure the total height to the roof peak (H), the height of the gutter or eave (G), the width (W), the length (L) and the width of the roof slope (S). The surface area is sum of the following: Area of side walls = 2 × (L × G) Area of sloping roof = 2 × (L × S) Area of end walls = 2 × [(G × W) + (0.5 × (H - G)) × W]

Total surface area = Area of side walls + area of roof + area of end walls Note that a multispan greenhouse only has 2 side walls, but every bay has roof areas and end walls. Tunnel house For a tunnel house, you need to measure the height (H), the width (W), the length (L) and the length of the curved frame (C). The surface area is sum of the following: Area of curved roof and walls = L × C Area of end walls = 2 × Π × (h × w) where Π = 3.14 Total surface area = Area of curved roof and walls + area of end walls

Straight walled curved roof For a straight walled curved roof greenhouse, you need to measure the height of the curved section (H), the width (W), the length (L) and the length of the curved frame (C). You also need the height of the vertical section of the side (S). The surface area is the sum of the following: Area of curved roof and walls = L × C Area of curved part of end walls = 2 × Π × (h × w) where Π = 3.14 Area of rectangular side walls = 2 × L × S Area of rectangular part of end walls = 2 × W × S Total surface area = Area of curved roof and walls + area of curved part of end walls + area of rectangular side walls + area of rectangular part of end walls

Note that a multispan greenhouse only has 2 side walls, but every bay has roof areas and end walls. Temperature difference (A) The temperature difference (A), called “delta T”, is the difference between the minimum required temperature in the greenhouse and the lowest outside temperature. The required temperature in the greenhouse is the set point that you want to achieve. This may be a compromise temperature to reduce heating costs. For example, while you may want a minimum night temperature of 19°C, to reduce your heating costs, you might decide to use a set point of 17°C instead. If the lowest outside temperature is 1°C and your set point is 17°C, the AT is 16 degrees. Calculating heat load due conduction The heat load due to conduction (QC) of your greenhouse equals the heat loss (U) multiplied by surface area (SA) multiplied by the temperature difference, divided by 1000. This will give you a heat load in kilowatts (kW): Qc = (U × SA × ΔT) / 1000 Additional accuracy in calculating the heat load can be achieved by including; The effect of wind Wind increases the removal of heat from the outside of the greenhouse so more heat can be transferred faster. If you are in a particularly windy area, especially during winter, it is a good idea to include it. The stronger the wind is, the greater the heat loss. A wind factor (W) is used in the heat loss calculation. Qc = (U × SA × ΔT × W) / 1000 The amount of heat loss through the floor The greenhouse floor is another surface where some heat can be lost. To include the loss of heat through the floor, you will need a U value for the floor (eg poured concrete is about 1.1, black plastic is about 2.7) and the surface area of the floor.

HEAT LOAD DUE TO LEAKAGE (QL) Air leakage is usually the second biggest source of heat loss from a greenhouse. To include the heat load due to air leakage (QL), you will need to know the air volume of the greenhouse in cubic metres (V), the number of air changes occurring (E) and the wind factor (W) and the difference between your temperature set point and the outside minimum temperatures (ΔT). Leakage (QL) = 0.373 × ΔT × V × E × W For the wind factor (W), refer to the table below: Wind Speed (km/hr)

Wind factor (W)

< 25

1.0

30

1.025

35

1.05

40

1.075

CALCULATING GREENHOUSE AIR VOLUME To calculate greenhouse air volume (V), imagine the structure as different shapes (rectangular, triangular or half cylindrical) and use the following formulae. Volume of a rectangular sectionFormula: Volume = Length × Width × Height Volume of a gable sectionFormula: Volume = 0.5 × Length × Width × Height Volume of a tunnel section Formula: Volume = Length × 6.28 × Width × Height If the greenhouse is made up of more than one shape, work out the volume for each and add them together. All dimensions should be in metres to give you a volume in cubic metres. For the number of air changes (E), use the following table. Greenhouse design

Air exchanges (E) (Leakage)

Single polythene film and metal frame

1.0

Double polythene film and metal frame

0.7

Single layer of glass and metal frame

1.08

Heater capacity Once you know what the maximum heat load of your greenhouse is, you can work out what capacity of heating system that you need. This is dependent on how efficient the heater is. In general, using bottled or natural gas, the heating efficiency is about 80%. Heater capacity = Heat load of your greenhouse (Q) / Heater efficiency Reducing heating requirements There are a number of strategies that can be used to reduce heating costs. •

•

• • • • • •

Use a greenhouse design that minimises surface area relative to production area, for example, a gutter connected multispan greenhouse has a smaller surface area than a number of separate greenhouses of the same total production area. Use cladding materials that have a low U value, for example, a double skin polythene clad greenhouse loses less heat than a single skin polythene covered greenhouse. Use thermal screens which have a lower U value than the cladding material. Close air leaks and repair any damage in the cladding materials. Make sure doors and vents close tightly. Use windbreaks to reduce the speed of wind passing over the greenhouse. Use the most efficient heater possible. Use an automated control system. Use cooler set points when possible, for example when controlling to a 24 hour average temperature regime, increase the day temperature and lower the night temperature.

EVAPORATIVE COOLING Evaporative cooling uses the natural relationship between relative humidity, water and air temperature. When water is evaporated it has a

cooling effect. Humidity is also increased and the vapour pressure deficit is reduced. Evaporative cooling is most effective in southern coastal areas and most inland areas.

Air temperature can be measured as either a dry bulb temperature or a wet bulb. The wet bulb temperature, gives an indication of what temperature air can be cooled to with evaporative cooling. Wet bulb temperature is measured using a thermometer with a wet sleeve or wick around it. Air passing over the thermometer and the water on the sleeve evaporates and cools the thermometer, so that it is reading less than the temperature read by a normal (dry) thermometer. The amount of cooling that can be achieved from evaporative cooling systems is dependent on how much water can be evaporated so is related the amount of water already in the air. This is relative humidity. Table 1.1 shows the temperature to which air can be potentially cooled. As can be seen from the table, evaporative cooling is most effective when the relative humidity is below 60%. Fan and pad systems Fan and pad systems combine two pieces of equipment. An exhaust fan is located at one end of the greenhouse and a porous pad is built into the wall of the structure at the opposite end. A pump circulates water over and through the pad. When the fan is in operation, it pulls air from outside the structure, through the evaporative pad, into the greenhouse. The air, passing through and over the wet pad evaporates some of the water and is cooled. As a result, cool air is drawn into the greenhouse to replace the hot air expelled by the fan.

Table 1.1 Potential cooling effect at different levels of relative humidity

These systems are quite effective for cooling but are relatively expensive to install and maintain. A disadvantage of the fan and pad system is that it tends to create a significant temperature gradient from one end of the greenhouse to the other (warmest at the fan end and coolest at the pad end) which can affect crop uniformity and make management more difficult. Under extreme conditions which demand a lot of cooling, the air movement through the greenhouse generated by exhaust fans may damage plants. Fogging systems Fogging systems are a fairly effective and uniform method of greenhouse cooling and provide a reasonable increase in relative humidity in a greenhouse. On a hot day, a cooling effect of up to 10oC can be achieved. Fogging systems produce very small droplets of water in the range of 10-20 microns which are suspended in the air and evaporate before they have time to fall on to the crop canopy. Fogging line pressure and fogging nozzles need to be properly maintained. A poorly maintained system may not provide sufficient cooling or could result in wetting of leaves and fruit. This can lead to reduced product quality and an increased incidence of pests and diseases. Misting systems Misting systems can be used to deliver a fine spray of water into the greenhouse air space to provide some relief during very hot and/or dry conditions. Most misting systems operate at a pressure of between around 250kPa and 300kPa. The water droplets are typically in the range of 100 to 200 microns. This sized droplet is too large to be completely evaporated and

so it falls quickly, wetting the crop and the greenhouse floor. While misting systems can have a cooling benefit, a wet canopy can lead to an increase in diseases and fruit damage. A wet floor can pose a safety risk to people in the greenhouse.

GREENHOUSE SCREENS There are a number of different materials in this category. In broad terms they can be divided into thermal screens, insect screens and shade screens. Thermal screens will probably be the most useful piece of equipment after the greenhouse itself, the hydroponic system, the heating system and the automated control system. Thermal screens Thermal screens offer a flexible and efficient way of improving the management of the greenhouse environment. These screens are designed to keep heat and radiation in a greenhouse during cold periods such as night time and prevent radiation and associated heat entering the greenhouse during hot periods, such as in the middle of a summer day.

Thermal screens offer a flexible and efficient way of managing the greenhouse environment

This dual capacity makes these materials suitable to a range of conditions throughout the year. They are moveable so that conditions can be adjusted and optimised through out the day and night. Thermal screens can reduce temperatures during the day by as much as 10°C and maintain temperatures during the night by as much as 5°C. When heating, thermal screens trap warm air nearer the crop, reduce the volume of air space to be heated and reflect radiation back to the crop. This

can reduce heating costs significantly - possibly by up to half. Under extreme, hot conditions, thermal screens are also valuable in reducing incoming radiation during the day. This reduces the heat load in the greenhouse and assists in maintaining humidity around the plants and reducing plant stress. The use of screens needs to be carefully managed to make sure that plants remain actively growing. Shade screens Shade cloth provides some of the functions of thermal screens. Under extreme, hot conditions, shading reduces the amount of incoming radiation. This in turn reduces the heat load in the greenhouse and assists in maintaining humidity around the plants and reducing plant stress. The use of shade cloth for cooling is most effective when the material is used outside of the greenhouse. If shade screens are used, pale coloured materials should be selected as these uniformly reflect solar radiation and do not absorb as much heat themselves as darker materials. A range of products offer shading from 30% up to almost total blackout. Whitewash paints are another shading option that can be applied to reduce the amount of radiation entering the greenhouse. White-wash is a lime wash or diluted paint applied to the outside of the greenhouse covering. It reduces the amount of solar radiation entering the greenhouse and therefore reduces temperature. The disadvantage of this type of screen is that it is applied seasonally, such as at the beginning of summer and consequently is not be readily adjusted for changing conditions from day to day or during a day. This results in reduced light levels even when temperature reduction is not needed. The end effect can be quite significant reductions in light levels and consequently yield. There are white wash products available which become transparent when wet so as to allow more light in during rain periods. During warm overcast conditions, these products can be hosed down to increase light transmission. Insect screens Insect screens are used to exclude flying and wind-borne pests from the greenhouse.

Insect screens keep flying and wind borne pests out of the greenhouse

The main disadvantage of insect screens is that they restrict airflow. This can have a significant impact on venting capacity of a structure. The size of the hole is the key characteristic that determines whether or not it can prevent a pest from getting into the structure. However, the finer the screen, that is, the smaller the hole size, the less air that can flow through it. Light in greenhouses Light is measured differently depending on what part of the light spectrum is being measured. The total light spectrum coming from the sun (400 to 1100 nanometer wavelengths) is measured in units of watts/m2. On a clear sunny summer day, there may be 1000 Watts/m2. Light may be measured in terms of its intensity (lux) or the number of photons reaching a surface (photon flux density). The part of the spectrum that humans can see, called visible light (380 to 770 nanometer wavelengths) is measured in lumens. The lumen is the metric unit of light intensity and the term lux refers to the number of lumens per square metre of surface area. In horticulture, the number of photons reaching a surface is more important. Photons are basically packets of energy which make up a stream of light. The number of photons trapped by a leaf determines the level of photosynthesis and therefore the amount of plant growth. The part of the spectrum that plants use is called photosynthetically active radiation (PAR) and relates to light in the 400 to 700 nanometer wavelength which is almost the same as visible light but not quite. It is measured in units of µmol/ m2/s and describes the photon flux density, that is, the number of packets of energy which reach a surface. Light Transmission The amount of light entering a greenhouse is influenced by;

• the orientation of the structure • the materials used in construction and covers • the shape of the roof. The greenhouse should be positioned north-south to provide more uniform light and reduce the shading effect of the support structure. The support structure must also be minimised to avoid shading. Metals make good structural material because of their strength which means narrower trusses and purlins can be used. A typical greenhouse frame can reduce light transmission by more than 10%. The type of covering material will also influence the level of light in the greenhouse. Finally the shape of the roof will impact on how much light enters the greenhouse. For example, a flat roof will limit the amount of light due to reflection while a curved roof provides the greatest annual light transmission. Quality of Light A balance of light across the PAR range is considered to be preferable, however there is increasing research being conducted in the area of light spectrum modification for improved plant growth. Diffuse light is better than direct light because it is able to reach the lower parts of the canopy (less shadowing) and it will not cause sunburn. Irrespective of whether the light is direct or diffuse, it must be of sufficient intensity (lux). The selected covering material may also be used to increase the amount of diffuse light. A textured surface on glass, for example, can increase the proportion of diffuse light without significantly reducing the total level of light transmitted. Coloured films The colour of plastic films affects the total level of light that enters the greenhouse. A clear film will transmit the most amount of light. Blue and green coloured plastics will transmit a lot of the light in the blue to blue-green wavelengths, but cut out much of the light in the red wavelengths. From the diagram above looking at PAR, it can be seen that red light is the most efficient waveband for plant growth. A blue plastic is likely to produce a

slower growing, shorter, tougher plant. Also of interest is that plants have been shown to use far-red light as a way of determining how much competition there is for light. This is because green surfaces, such as leaves from other plants, reflect a lot of far-red light. If the plant perceives that there is a lot of competition, it will put less energy into growing roots and more into growing tall, quickly. A green plastic is likely to produce a stretched, slow growing, poor performing plant. A white film will reduce the total amount of light transmitted by as much as 20%, but the light spectrum entering the greenhouse will remain similar to the natural light spectrum. Light Intensity Plants have an optimal intensity of light. This is the point at which the process of photosynthesis is maximised and plant growth is greatest. If the level of light is less, growth is reduced. The point where an increase in light intensity will not increase photosynthesis any more is called light saturation. Cucumbers for example require at least 250 µmol/m2/s of photosynthetically active radiation (PAR). Below this level, productivity will decline. The amount of light in the greenhouse should be maximised wherever possible. The management of heat is the only reason for reducing incoming light levels. What to aim for in the greenhouse Low light slows growth and increases the cost of production but excessive light intensity can damage some plants and/or fruit. Light is increased by minimising objects above the plants including frames, pipes, lights and other equipment. The level of radiation entering a greenhouse can be reduced with screening materials. Supplementary lighting Supplementary lighting is not considered to be economical for producing most crops in Australia, however, supplementary light may be used to improve seedling uniformity during propagation.

GREENHOUSE COVERING MATERIALS

The covering material used on a greenhouse influences the productivity and performance of a structure. Covering materials impact on the level and quality of light available to the crop. Diffused light is better than direct light. Fluorescent and pigmented films can increase the proportion of good red light. Dust, attracted to plastic films, will reduce the transmission of radiation. Water droplets on the inside of coverings have been shown to reduce light transmission by 8% and will also block thermal radiation. Greenhouse coverings all reduce light to some extent. As coverings become dirty and as they get older, less light enters the greenhouse. Condensation (water drops) on the covering material also reduce light. Light coloured materials in the greenhouse, such as white weed matting, increase the light available to the crop. Key characteristics that should be considered in selecting a covering material are the cost, its durability (how long it lasts), its weight and ease of repair or replacement, how much light is transmitted through the material and how much energy moves through the material. Diffusing materials are designed to scatter incoming light and result in better light conditions for crops – for example, a cloudy white plastic film diffuses light better than a clear plastic film. Glass Glass has long been the traditional covering. Its favourable properties include: • • • • •

high transmission in the photosynthetically active radiation (PAR) bandwidth good heat retention at night low transmission of UV light durability low maintenance costs.

Plastic Sheeting Essentially there are three materials in this category - polycarbonate, acrylic (polymethyl methacrylate) and fibreglass. Plastic sheeting is not used extensively in Australia but its use is increasing. Sheeting products are more durable than plastic films and have fairly good heat retention, good initial

transmission in the PAR range and low UV light transmission. Plastic films Films are the most common and lowest cost type of covering material. The types of film available are polythene (polyethylene), EVA (ethyl vinyl acetate) and PVC (poly vinyl chloride). With the constant improvements in plastics, these covering materials offer a lot of flexibility and performance options. Coverings can have a variety of additives which are used to give plastic films useful properties. For example, films may be used to exclude ultra violet (UV) light for chemical free pest control or reflect long wave infra red (IR) radiation to improve heat retention at night. As a result, some plastic covering materials are coloured or tinted. Additives to the plastic determine its; • • • • •

durability capacity to reduce heat loss capacity to reduce droplet formation transmission of particular wavelengths of light capacity to reduce the amount of dust sticking to the film.

Types of Additives 1. UV (290-400 nm) absorbers and stabilisers increase durability, reduce the potential damage to biological systems in the greenhouse and may control some plant pathogens 2.

Infrared (700-2500 nm) absorbers reduce long wave radiation and minimise heat loss

3. Long wave radiation (2500-40000 nm) absorbers reduce the loss of heat radiated from materials and objects (including plants) inside the greenhouse 4. Light diffusers scatter light entering the greenhouse, reducing the risk of plants getting burnt and improving the amount of light available to the lower parts of the plant 5. Surfactants reduce the surface tension of water, dispersing condensation 6.

Antistatic agents reduce the tendency of dust to accumulate on plastic films.

In addition, 1. Colour pigments may improve plant growth by altering the proportion of selected wavelength ranges 2. Fluorescence may be used to increase the emission of red light 3. Glossy surfaces may repel insects The process of making multilayer films enables thin layers of materials with different properties to be joined to make superior composite films. Properties such as durability, creep (deformation over time) and long wave radiation absorption can be improved. Maintenance A poorly maintained covering material can lose a lot of energy and significantly increase production costs. Glass coverings should be kept clean and broken panes replaced. Plastic coverings need to be replaced routinely. The performance of plastic coverings declines over time. Old coverings reduce light transmission which can restrict yield. The useful life of plastic films depends on the specifications of the plastic purchased. All plastic covering materials need to be replaced before they visibly start to break down; discolouration, for instance, is an early indication of wearing.

UNIFORMITY IN THE GREENHOUSE Uniformity in the greenhouse is a basic requirement for achieving a good, high yielding uniform crop. Uniformity increases productivity, labour and resource efficiency and product quality. One of the main areas of variation in a greenhouse is the irrigation system. A high distribution uniformity is critical. Temperature and humidity can also vary significantly in the greenhouse and affect crop growth and production. In some situations, carbon dioxide levels become depleted and affect crop growth. There are three key areas that need to be addressed are to make sure the greenhouse environment is as uniform as possible.

Ventilation Adequate ventilation is necessary to avoid the build up of excess heat, humidity and depletion of carbon dioxide. Both passive (using vents) and active (using fans) ventilation should be capable of completely exchanging the air in the greenhouse at least once every minute. It is better to have more venting capacity than you use, rather than not enough. Roof ventilation is also better than side wall ventilation. Vents in the walls of a greenhouse are limited in their capacity to provide an even environment. While they are very effective at exchanging air in the first couple of rows of the crop, the crop itself acts as a windbreak. This results in very little air movement towards the middle of the crop. Air circulation Moving and mixing of air throughout the greenhouse evens out temperature and humidity differences. Without sufficient air circulation, pockets of hot and cold air can occur. Humidity will also build up excessively around the plants. Side vents are limited in their capacity to fully mix the air in a greenhouse (a factor of the span width of the greenhouse). Roof ventilation circulates air well when in use. Fans can be used to improve air circulation and reduce uneven growing conditions. Heating Heating is relatively expensive and needs to be efficient. A heating system must be able to distribute heat evenly throughout the crop. Particular parts of the greenhouse will be hotter or colder. The northern end and western side of a structure will tend to be hotter. The southern end will be colder. The northern end will also have less shading. Screens can be used to reduce the extremes and improve uniformity. Light in Greenhouses Light is measured differently depending on what part of the light spectrum is being measured. The total light spectrum coming from the sun (400 to 1100 nanometer wavelengths) is measured in units of watts/m2. On a clear sunny summer day, there may be 1000 Watts/m2. Light may be measured in terms of its intensity (lux) or the number of photons reaching a surface (photon flux density). The part of the spectrum

that humans can see, called visible light (380 to 770 nanometer wavelengths) is measured in lumens. The lumen is the metric unit of light intensity and the term lux refers to the number of lumens per square metre of surface area. In horticulture, the number of photons reaching a surface is more important. Photons are basically packets of energy which make up a stream of light. The number of photons trapped by a leaf determines the level of photosynthesis and therefore the amount of plant growth. The part of the spectrum that plants use is called photosynthetically active radiation (PAR) and relates to light in the 400 to 700 nanometer wavelength which is almost the same as visible light but not quite. It is measured in units of µmol/ m2/s and describes the photon flux density, that is, the number of packets of energy which reach a surface. Light Transmission The amount of light entering a greenhouse is influenced by; • the orientation of the structure • the materials used in construction and covers • the shape of the roof. The greenhouse should be positioned north-south to provide more uniform light and reduce the shading effect of the support structure. The support structure must also be minimised to avoid shading. Metals make good structural material because of their strength which means narrower trusses and purlins can be used. A typical greenhouse frame can reduce light transmission by more than 10%. The type of covering material will also influence the level of light in the greenhouse. Finally the shape of the roof will impact on how much light enters the greenhouse. For example, a flat roof will limit the amount of light due to reflection while a curved roof provides the greatest annual light transmission. Quality of Light A balance of light across the PAR range is considered to be preferable, however there is increasing research being conducted in the area of light spectrum modification for improved plant growth. Diffuse light is better than direct light because it is able to reach the lower parts of the canopy (less shadowing) and it will not cause sunburn.

Irrespective of whether the light is direct or diffuse, it must be of sufficient intensity (lux). The selected covering material may also be used to increase the amount of diffuse light. A textured surface on glass, for example, can increase the proportion of diffuse light without significantly reducing the total level of light transmitted. Coloured films The colour of plastic films affects the total level of light that enters the greenhouse. A clear film will transmit the most amount of light. Blue and green coloured plastics will transmit a lot of the light in the blue to blue-green wavelengths, but cut out much of the light in the red wavelengths. From the diagram above looking at PAR, it can be seen that red light is the most efficient waveband for plant growth. A blue plastic is likely to produce a slower growing, shorter, tougher plant. Also of interest is that plants have been shown to use far-red light as a way of determining how much competition there is for light. This is because green surfaces, such as leaves from other plants, reflect a lot of far-red light. If the plant perceives that there is a lot of competition, it will put less energy into growing roots and more into growing tall, quickly. A green plastic is likely to produce a stretched, slow growing, poor performing plant. A white film will reduce the total amount of light transmitted by as much as 20%, but the light spectrum entering the greenhouse will remain similar to the natural light spectrum. Light Intensity Plants have an optimal intensity of light. This is the point at which the process of photosynthesis is maximised and plant growth is greatest. If the level of light is less, growth is reduced. The point where an increase in light intensity will not increase photosynthesis any more is called light saturation. Cucumbers for example require at least 250 µmol/m2/s of photosynthetically active radiation (PAR). Below this level, productivity will decline. The amount of light in the greenhouse should be maximised

wherever possible. The management of heat is the only reason for reducing incoming light levels. What to aim for in the greenhouse Low light slows growth and increases the cost of production but excessive light intensity can damage some plants and/or fruit. Light is increased by minimising objects above the plants including frames, pipes, lights and other equipment. The level of radiation entering a greenhouse can be reduced with screening materials. Supplementary lighting Supplementary lighting is not considered to be economical for producing most crops in Australia, however, supplementary light may be used to improve seedling uniformity during propagation.

2: IPM Scouting and Decision-making INTRODUCTION Integrated pest management (IPM), also known as integrated pest control (IPC) is a broad-based approach that integrates practices for economic control of pests. IPM aims to suppress pest populations below the economic injury level (EIL). The UN’s Food and Agriculture Organisation defines IPM as “the careful consideration of all available pest control techniques and subsequent integration of appropriate measures that discourage the development of pest populations and keep pesticides and other interventions to levels that are economically justified and reduce or minimize risks to human health and the environment. IPM emphasizes the growth of a healthy crop with the least possible disruption to agro-ecosystems and encourages natural pest control mechanisms.” Entomologists and ecologistshave urged the adoption of IPM pest control since the 1970s. IPM allows for safer pest control. This includes managing insects, plant pathogens and weeds. Globalization and increased mobility often allow increasing numbers of invasive species to cross national borders. IPM poses the least risks while maximizing benefits and reducing costs.Integrated Pest Management (IPM) is the coordinated use of pest and environmental information along with available pest control methods, including cultural, biological, genetic and chemical methods, to prevent unacceptable levels of pest damage by the most economical means and with the least possible hazard to people, property, and the environment”. Integrated means that all feasible types of control strategies are considered and combined as appropriate to solve a pest problem. Pests are unwanted organisms that are a nuisance to man or domestic animals, and can cause injury to humans, animals, plants, and property. Pests reduce yield and/or quality in plants ranging from field crops, fruits and

vegetables, to lawns, trees, and golf courses. Management is the process of making decisions in a systematic way to keep pests from reaching intolerable levels. Small populations of pests can often be tolerated; total eradication is often not necessary, or feasible.

THE BASICS OF IPM All of the components of an IPM approach can be grouped into three activities. The first is monitoring; the second is assessing the pest situation; and the third is taking action. Trace these steps through this web site by reading through these pages. For more information follow the links on each page. IPM is information intensive and relies on scouting and monitoring programs for the collection of field data about key factors such as: • • • • • •

Pest population identification Disease pressure Weather conditions and degree-days Pest date of first occurrence of biological events in their annual cycle Crop growth stage Presence, reliance and preservation of beneficial organisms IPM uses decision support systems for determining if control measures are necessary and what measures are most appropriate. Such as: • • • • •

Economic thresholds - the pest population level that inflicts crop damage greater than the cost of control Availability of selective pesticides Action levels - pest level when action should be applied to prevent pest from reaching injurious levels Environmental risk measurements (i.e. impacts on pollinators) Disease forecasting systems IPM programs seek to avoid pest damage through practices such as:

• Use of field sanitation and reduction of pest habitat • Crop rotations

• Selection of pest/disease tolerant or resistant seeds and varieties • Judicious use of pesticides that prevent pest infestations • Resistance management Why Practice IPM? You might be wondering why you should even consider IPM when pesticides so often succeed at controlling pests. Here are some reasons for using a broader approach to pest management than just the use of pesticides. • Many IPM practices are used before a pest problem develops to prevent or hinder the buildup of pests. • Keep a Balanced Ecosystem. Every ecosystem, made up of living things and their non-living environment, has a balance; the actions of one creature in the ecosystem usually affect other, different organisms. Many of our actions in an ecosystem can change this balance, destroying certain species and allowing other species (sometimes pests themselves) to dominate. Beneficial insects, such as the ladybird beetle and lacewing larvae, both of which consume pests, can be killed by pesticides, leaving fewer natural mechanisms of pest control. • Reliance on Pesticides can be Problematic. Pesticides are not always effective when used as a singular control tactic. Pests can become resistant to pesticides. In fact, some 600 cases of pests developing pesticide resistance have been documented to date, including populations of common lamb-quarters, house flies, Colorado potato beetle, Indian meal moth, Norway rats, and greenhouse whitefly. • IPM Is Not Difficult. You will have done much of the “work” for an IPM approach if you’ve figured out the problem (the pest), determined the extent of the pest population, and decided on the best combination of actions to take. • Maximize Effectiveness of Control Tactics. Pest control practitioners, following traditional programs, sometimes apply pesticide treatments on a calendar based schedule regardless of the stage of development of the target pest and the number of pests present. Using an IPM approach will ensure that all control tactics, including pesticides, are used at the proper time and only to reduce pest damage to acceptable levels. This will reduce costs from unnecessary pesticide applications and insure that control tactics are used when they will be most effective.

•

Promote a Healthy Environment. The definition of IPM promotes a careful consideration of all pest control options with protection of the environment a key goal. • Natural Enemies Conserved. Parasites and predators are part of the natural control mechanism for some pest populations. These natural controls are considered and protected in an IPM program • Maintain a Good Public Image. A thoughtful approach to pest control, which protects the environment and provides an abundant, affordable crop and safe living conditions, is a basic goal of IPM. A regular monitoring program is the basis of IPM decision making, regardless of the control strategies used. By regular monitoring, a scout is able to gather current information on the identity and location of pest problems and to evaluate treatment effectiveness. The following are the basics of scouting programs in New England with growers who participate in Greenhouse IPM Programs.

TOOLS USED IN GREENHOUSE IPM The list of essential monitoring tools includes: • Trained personnel • Hand-lens with 10x power and/or optivisor (headset with magnifying glass) • Yellow sticky cards, clothes pins, bamboo stakes • Flagging tape or colored flags • Record-keeping system (clipboard or small notebook and pen) • Individual maps of all greenhouses • Support labs for disease diagnosis and soil tests and/or solubridge if a soil-testing laboratory is not available • Resource information such as pesticide labels, pictures and life cycles of key pests and “common sense”. Additional monitoring tools: • Soil thermometer • Field microscope • Potato slices (knife, potatoes) to monitor fungus gnat larvae

• Waterproof marker to number sticky cards. Pre-Crop Site Evaluation One month prior to the introduction of a crop, evaluate the entire greenhouse, inside and out. Note the presence of weeds in and around the greenhouse, drainage problems, algae build-up, pet plants, overwintered plants such as impatiens or geraniums and debris under benches. Crops growing in adjacent greenhouses or outdoors should be recorded. Previous pest problems in the greenhouse and current pesticide application methods should be reviewed. A plan of action may then be developed to eliminate these problems prior to the arrival of the crop. Prevention of key pest problems may be more easily accomplished if the grower and scout take the time to identify, analyze and correct problems before crops are introduced. Also, consider how the variety of plants to be grown in the same area may influence ease of pesticide applications and spread of disease. For example, keep seedling and cutting geraniums separate to help minimize spreading bacterial blight. Keep propagation houses separate from other growing areas, and vegetable transplants separate from ornamentals to help reduce the incidence of Impatiens Necrotic Spot Virus when Western Flower Thrips are present. Also, most pesticides labeled for ornamentals are not labeled for vegetable crops. Inspection of Incoming Plants At the time of arrival or soon after, the scout should inspect one-third or more of the plants. Thoroughly examine the plants for signs of insects and diseases (see chart). Early detection and prompt action can minimize the spread of insects and diseases and save pesticide applications.

WEEDS, ALGAE, AND LIVERWORTS Weed Management Inside The Greenhouse Maintaining weed-free growing conditions is an essential part of producing high-quality greenhouse crops. Insects and diseases can be kept to a minimum only if proper weed control practices are carried out regularly, along with other appropriate controls. Weeds may compete with crops for light, water and nutrients; reduce the

aesthetic value of crops; and create a poor impression. Weeds are also a primary source of aphids, whiteflies, leafminers, thrips, mites, slugs and diseases. Low-growing weeds help maintain moist conditions, which favor fungus gnats and shore flies. Many common greenhouse weeds such as chickweed (Stellaria media), oxalis or woodsorrel (Oxalis spp.), bittercress (Cardamine hirsuta), jewelweed (Impatiens spp.), dandelion (Taraxacum officinale) and ground ivy (Glechoma hederacea) can host tospoviruses including impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV) while showing few, if any, symptoms. Thrips can vector these viruses to susceptible greenhouse crops. Weeds can also carry other plantdamaging, aphid-vectored viruses. An integrated weed management program can help manage weed populations. This approach includes preventive measures such as sanitation and physical barriers, and control measures such as hand weeding and the selective use of postemergence herbicides. Prevention Weed seeds are easily blown into the greenhouse though vents and other openings. Weeds and their seeds can enter a greenhouse on plants, tools and equipment. Seeds can be moved in soil and by wind, irrigation water, animals and people. Creeping wood sorrel, (Oxalis corniculata), hairy bitter cress (Cardamine hirsuta), prostrate spurge (Euphorbia humistrata), common chickweed (Stellaria media) and other weeds are persistent problems in greenhouses, reproducing primarily by seed, with several generations each year. Prevention is the grower’s first line of defense. Sanitation Keep weed seeds, rhizomes and other propagules out of the greenhouse by using sterile media and “clean” plant materials, and by controlling weeds outside the greenhouse. Clean up spilled growing media inside and outside the greenhouse.

Inspection of Incoming Plants Key pest

How to monitor

Where to look

Melon Aphid (Aphis gossypii)

Rely on plant inspection, not sticky cards. Scout weekly, early in the crop, before flowering. Look for small, 1/16

Inspect incoming plant material, on underside of leaves and stems. Melon aphids are more likely to

inch long aphids with dark cornicles or “tailpipes.” Melon aphids are less likely to form winged adults than green peach aphids.

be found along the plant stem than on the growing tip.

Green Peach Aphid (Myzus persicae)

Monitor weekly. Rely on plant inspection, not sticky cards. Winged adults are found on cards when aphid colonies on weeds and crops become overcrowded.

Look on tips of new growth for 1/14 inch long green to pinkish aphids. Look for signs of aphid activity: shed white skins, honeydew, and presence of ants. Inspect and remove weeds.

Western flower thrips (Frankliniella occidentalis)

Rely on sticky card counts for population trends and to evaluate treatments. Use cards at floor level to detect overwintering thrips, beginning in February. Place cards at bench level, just above crop in March, before plant damage occurs (April - September). Cards placed at HB level, and in mist propagation areas will detect fewer thrips.

Inspect incoming plant material for adults and larvae by tapping tender new growth and flowers over a white sheet of paper. Keep plants isolated for 4-5 days to detect emerging eggs and pupae. Inspect and control weeds outside of the greenhouse in early spring, especially white clover.

Whiteflies (Bemisia tabaci, Trialeurodes vaporariorum)

Rely on plant inspection to detect immature stages, especially on cuttings and young plants. If using insect growth regulators, use indicator plants to assess treatment effectiveness. Use sticky cards to monitor adults. Sequential sampling is an effective time-saving method for poinsettias. Place card horizontally if usingEncarsia formosa for biological control.

Older (3rd and 4th instar) immatures are found on the lowermost leaves. Egg-laying adults are found on the uppermost leaves. Inspect and remove weeds and “pet plants.”

Fungus gnats & shoreflies (Bradysia sp.)

Use sticky cards to monitor for adults. Place cards just above soil surface. Horizontal placement will attract more adults. Use potato slices (11/4” long by

A high emergence of adults may occur after watering dry pots. Favorable habitats include areas with standing pools of water, muddy floors, and weeds.

1”) to monitor for larvaeespecially during cool, moist weather. Examine daily. Pythium root and stem rots (Pythium sp.)

Visually examine roots for cortex that “sloughs off” leaving central core on geraniums, impatiens, snapdragons, vinca, poinsettias etc. Stem cankers are brown to black. Monitor fertility and EC levels.

Monitor incoming plants and plants that may have been stressed by high salt levels, wounding, and transplant shock, especially if fungus gnats or shore flies are present.

Rhizoctonia damping off, root rot, stem

Monitor seed flats of susceptible plants such as begonia, impatiens, petunia, dahlia for post-emergence damping off.

Monitor seed flats for damping off especially near walkways. Web blight may occur when

canker and web blight (Rhizoctonia solani)

Look for small water-soaked spots on stem or leaves before seedlings collapse. Look for cobwebby growth that mats leaves together (web blight).

bedding plants are placed close together during humid, warm conditions.

Botrytis Blight (Botrytis cineraria)

Monitor closely during favorable conditions, ie. cool temperatures, free moisture and presence of fungal spores, fuzzy gray to brown. Flowers may fade early and then mat together.

Look on tender tissues (flowers, terminal buds or cuttings, or weakened tips of leaves for soft, tan to brown dead areas, and gray fungal growth. Monitor areas with poor air circulation, and crowded plants.

Powdery Mildew (Erysiphe sp., Oidium sp.)

Look for white powdery growth esp. on upper leaf surface of roses, begonias, viola, phlox, chrysanthemums. On poinsettia, look for white or yellow spots on upper leaf surface. White patches up to 1/2 inch in diameter may be on the lower or upper surface.

Monitor closely in areas with poor air circulation, high humidity or drafty places with more temperature fluctuations between day and night temperatures.

Bacterial Blight (Xanthomonas pelargonii)

Inspect geraniums more closely during warm weather. Look for isolated leaf wilting, V- or wedge-shaped yellowing between the veins and 1/8 round, brown spots. Look for vascular discoloration. Plants may wilt and die.

All geraniums are susceptible. Do not place ivy geraniums over geraniums. When infected, they often do not show any distinct symptoms, perhaps only loss of vigor, and will serve as innoculum source. Monitor areas closely with geraniums from different suppliers present. If possible, buy from one supplier.

When moist, it provides an ideal environment for the germination of weed seeds. Screen vents and other openings to help limit the entry of windblown seed as well as insects. When scouting, identify the type of weeds (broadleaf or grass), life cycle (annual, biennial or perennial) and location. It is critical to remove weeds from pots, benches and floors before they flower and produce seeds. For example, a single plant of bittercress can produce 5000 seeds that can germinate in as little as 5 days and can propel the seeds over 9 feet from the plant. Creeping woodsorrel also expels seeds by force throughout a greenhouse. Physical Barriers Use a physical barrier like weed block fabric to limit weed establishment on greenhouse floors. Leave the fabric bare so it can be easily swept.

Covering the weed fabric with gravel makes it difficult to remove spilled potting media, providing an ideal environment for weed growth. Regularly pull escaped weeds before they go to seed. Repair tears in the weed block fabric. Controlling Existing Weeds These methods may be used to control existing weeds: 1) hand pulling, and 2) a postemergence herbicide. These measures do not prevent reseeding of weeds. Precautions on Herbicide Use Few herbicides are labeled for use in a greenhouse due to the potential for severe crop injury or death to desirable plants. This injury may occur from 1) spray drift if fans are operating at the time of application, and 2) volatilization (changing from a liquid to a gas). Herbicide vapors are easily trapped in an enclosed greenhouse, and can injure desirable plant foliage. Use only herbicides labeled for use in the greenhouse. Carefully follow all label instructions and precautions. It is the applicator’s responsibility to read and follow all label directions. Use a dedicated sprayer that is clearly labeled for herbicide use only. Symptoms of Herbicide Injury Herbicide injury symptoms include discolored, thickened, or stunted leaves. Sometimes the growing point of young seedlings is injured, severely stunting growth. Symptoms may be similar to those of nutrient imbalances, viral diseases or air pollutants from a faulty heating system. Proper diagnosis is needed to determine the cause. Symptoms can be so severe that injured plants cannot be sold.

3: Identifying Pest and Beneficial Insects on Sticky Cards Yellow or blue colored sticky cards are available in various sizes. Blue cards may be more attractive to thrips while yellow cards are used for overall monitoring. Yellow sticky cards are used to detect infestations of adult flying insects in greenhouses (fungus gnats, leafminers, shore flies, western flower thrips, whiteflies, winged aphids). While sticky cards are available in different sizes, 3” by 5” sticky cards are most commonly used. To use sticky cards, attach each card to a wire or wood stake with a clothes pin. Another option is to glue two clothespins back-to-back. Attach one end of the clothespin to a stake and clip the card to the other clothespin. This will allow you to move the card upwards as the plant matures. For general monitoring, attach sticky cards vertically just above the plant canopy, about 4 inches. For fungus gnats, place cards horizontally just above the soil surface or lay them flat on rims of pots. Each yellow sticky card should be numbered and placed in the greenhouse at the minimum rate of one card per 1,000 sq. ft.. Space the cards equally throughout the entire range in a grid pattern. Place cards near all entryways and vents. Small greenhouses (< 4,000 sq.ft.) can be scouted as one unit. Larger greenhouses should be divided into 2,000 to 3,000 sq. ft. sections for ease of scouting. Change the cards weekly, and place new cards in the same areas of the greenhouse to track pest trends. Brief, concise and accurate information is one of the best tools available to make a pest management decision. Identify and record pest numbers in a notebook. Over time, population trends will emerge and provide direction for your pest management program.

INDICATOR PLANTS

Indicator plants are chosen from pest-infested plants in a greenhouse. The scout uses these plants to make a close, ongoing examination of a pest’s development through its life cycle and to monitor treatment effectiveness. Indicator plants should be marked and numbered with a colored flag or flagging tape so that the scout can identify them quickly each week.

MAKING PEST MANAGEMENT DECISIONS Each week, the grower and scout should review the scouting information. Pest numbers recorded from sticky card counts and foliar inspections, the use of indicator plants, and located reservoirs of pests will help to prioritize a pest-management strategy.

SCOUTING METHODS

Scouting •

Use yellow sticky cards to trap adult whiteflies, fungus gnats, winged aphids, leafminers, & shoreflies

Magnification Needed •

Use a 10x-20x handlens to see identifying characteristics of insects on sticky cards.

Pest Insects Trapped on Sticky Cards

• • • • • • •

Aphids Fungus Gnats Shore Flies (nuisance pest) Leafminers Leafhoppers Thrips Whiteflies

Winged Aphids • • • • • •

Aphids have pear shaped bodies with two cornicles or “tailpipes” at theirrear Legs & antennae are long and thin Trapped aphids may give birth to several nymphs before they die Wings tend to be spread on either side of their body on the sticky cards Wings are longer than their body Look for two parallel veins close to the edge with a darkened area.

Fungus Gnat Adults • • • •

Small, dark mosquito-like flies with grayish wings Have long, slender legs and antennae Look for distinct Y-shaped vein at the tip of the single pair of wings Bodies may be hump backed (depends upon species)

Shore Flies • Look for – three to five pale spots on their grayish wings – short bristle- like antennae – and moderately long legs • Have robust, stout body compared to fungus gnats • About the size of fruit flies Leafminer Adults • Small, robust flies with noticeable yellow patch on their body • Have short antennae and two transparent wings

• Have a large cannon-shaped structure at the end of the abdomen that is used to puncture leaves and lay eggs

•

Often confused with shore flies (look for yellow on their body) plus plantdamage

Leafhoppers • • • • •

Slender insects with short bristle like antennae Wings are held roof like over the abdomen Wedge shaped, tapering to the rear No antennae visible Color vary depending upon species

Leafhopper Adult Thrips

Thrips

• Generally, the smallest insects you will see on the cards • Narrow and cigar shaped • Look for red eyes, short antennae fringed wings with hairs on end to distinguish from grains of peat moss Whiteflies • Look for whitish bloom whichtends to disappear after a fewdays • Whiteflies becomes orangish in color as they blend into the sticky material on the trap • Slightly larger than thrips

Whiteflies

Banded Winged Whiteflies •

Adult - Mature adult bandedwinged whiteflies have zig-zag bands across the front pair of wings. The hind pair of wings are unmarked. With the exception of the front banded wings this whitefly is very similar to greenhouse whitefly. • Egg - The eggs are about 0.12 mm long and 0.10 mm wide. Eggs are placed randomly or in circles on the leaf underside. Newly deposited eggs are pale yellow and turn pale pinkish just before hatching.

Banded Winged Whiteflies

•

Nymph- Young nymphs are 0.37 mm long, and as nymphal stages progress become just over one-half mm long. They are translucent white, with a yellow spot on each side of the abdomen. When the first instar nymph first settles down it begins to secrete a wax fringe that will become the side walls of the pupal case. As growth occurs the nymphal stages will secrete a marginal fringe of translucent setae, and the dorsal medial area of the integument becomes brown. • Pupa- The pupal case is just short of 1 mm long and 0.5 mm wide. The translucentmarginal setae are of two lengths and the marginal palisade of wax rods is verydistinct. The dorsal medial region is dark brown and uneven; the operculum isyellowish brown. Beneficial Insects Trapped on Cards • Parasitic Wasps (many different types) – Often attracted to yellow sticky cards • Hunter flies, Hover Flies and otherBeneficial Flies

Parasitic Wasps • Often Hymenoptera species • May be stout or slender • In comparison with flies, often have longer, elbowed antennae and bodies may be more pointed toward the rear

Aphidius colemanii

•

Many have clear wings with only onedistinct, angular vein along the front ofeach forewing

Encarsia formosa •

Commercially available parasitic wasps used to control whiteflies (especially greenhouse whiteflies ) • Small, parasitic wasp with black head and thorax and yellow abdomen • May look like tiny black dots on yellow card. Eretmocerus sp. •

Commercially available parasitic wasp used against whiteflies (especially sweet potato whiteflies) • Yellow or straw colored • With elbowed antennae. Shore Fly Parasitoid

Hexacola sp. is a parasitic wasp that lays it eggs into shorefly larvae.

Shore flies (Scatella stagnalis) and fungus gnats are often considered together as greenhouse pests, but they belong to two distinct groups of insects. Shore flies feed on algae and are found in areas where algae are growing. Adult shore flies are small, dark-grey flies (approx. 1/8 inch long), which slightly resemble a Drosophila fruit fly, with a robust body and short legs and antennae. They have five distinctive whitish spots on their grey wings. Their single pair of wings lacks the characteristic Y-shaped vein at the tip seen in fungus gnats, and the shore fly adult has short antennae. Synacra pauperi

Fungus gnats, Bradysia spp. are major insect pests in greenhouse crop production systems. The adults are a nuisance, and can carry spores of soil and foliar pathogens on their bodies. The larvae cause direct injury by feeding on plant roots, which reduces the plant’s ability to take-up water and nutrients. Also, the wounds created by larval feeding provide an entry site for secondary soil-borne pathogens. In addition, fungus gnat larvae have been shown to directly transmit soil-borne pathogens such as Pythium spp. to plants. • Naturally occurring parasite of fungus gnats • Adults are about the same size as fungus gnats • Look for narrowing between the head and thorax & between thorax and abdomen • Abdomen tapers to a sharp tip • Antennae are beaded & elbowed

• May be seen in unsprayed greenhouses Hunter Flies

Shiny wings without spots

Adult hunter flies are being found on yellow sticky cards, especially in greenhouses that are using biological controls. This small fly, native to the Mediterranean region, has been observed in greenhouses in the US and Canada since 2002. Adults perch on a leaf and wait for its smaller prey. The adult hunter fly preys on other flying insects, catching them in flight. It feeds on fungus gnats and shoreflies, but also on other flying insects such as leafminer and, to a lesser extent, whiteflies. The larvae live in the soil and are generalist predators on soil-dwelling organisms such as fungus gnat and shorefly larvae. Larvae grow for about two weeks and then pupate in the soil for two weeks. Growers may confuse hunter flies with shore flies. However, hunter flies (Coenosia attenuata) have clear wings with no spots and are about twice the size of shore flies. They are in the same family as the housefly, but smaller than the common housefly. Hunter flies are originally from Europe. They were first found in the

United States in October of 1999 at a commercial greenhouse in upstate New York. Adult females lay eggs in the soil that hatch in about 5 days. The hunter fly larvae seek other soil dwelling insects such as fungus gnat larvae as prey. Hunter flies are not commercially available. Males are a lighter gray than females. Wings are uniformly clear (unlike shore flies). Hunter flies prey on fungus gnats, shore flies, leafmining flies. Hover Flies Hoverflies belong to a large family of small to large flies. They are true flies or Diptera, with only one pair of wings in the Family Syrphidae. (Wasps and bees have two pairs.) Many hoverflies have spots, bands or stripes of yellow or brown against a dark-coloured background, sometimes with dense hair covering the body surface (emulating furry bumblebees). Their fast flight, ability to hover and, in some species, their size are astonishing characteristics. Some hoverflies are among the largest flies of Central Europe. Many species are very colorful. It is not always that easy to identify hoverflies. Some thick-headed flies and beeflies are similar and their dark coloration makes it hard to identify them correctly at a glance. Beeflies tend to be longer, hairy, have snouts and are a study in themselves! Hovering is a speciality although other flies can also hover—the head of the insect remains absolutely still whilst in flight. They may be seen “Nectaring” on many wild and garden flowers where they are amongst the most frequent of visitors. In Holland and Belgium alone over 300 species exist! In Britain about 270 species are known at present, but significant species and numbers can migrate like butterflies with a powerful flight such as Scaeva pyrastri. The Marmalade Fly Episyrphus balteatus is one of the most common hoverflies to be seen in the garden. The distinctive double stripes on the abdomen make it almost unmistakable.

Many are seen in the summer season in numbers while mixing with butterflies, bees, bumblebees and other flower dependent insects. Male hoverflies tend to emerge and mature first, earlier in the season to ensure reproduction is successful. Many species are useful to the gardener since their larvae eat pest aphids on garden plants and crops. The degree to which they contribute to pollination is also ironically poorly investigated but no doubt are important for carrots, onions and fruit trees.

MISCELLANEOUS INSECTS Midges Midges are a group of insects that include many kinds of small flies. They are found (seasonally or otherwise) on practically every land area outside permanently arid deserts and the frigid zones. The term “midge” does not define any particular taxonomic group, but includes species in several families of Nematoceran Diptera. Some midges, such as many Phlebotominae (sand fly) and Simuliidae(black fly), are vectors of various diseases. Many others play useful roles as prey items for insectivores, such as various frogs andswallows. Others are important as detritivores, participating in various nutrient cycles. The habits of midges vary greatly from species to species, though within any particular family midges commonly have similar ecological roles. Midges are sometimes used in biotic indexes of waterquality assessment. Their presence, when ephemeroptera (mayflies), plecoptera (stone flies), and trichoptera (caddis flies) are absent or rare,

generally is considered an indication of poor water quality. The tiny wormlike aquatic larvae, soft-bodied and often bloodred, are commonly known as bloodworms. They are important food for aquatic animals, especially trout and young salmon. The nonbiting midge is related to the biting midge, which is in the family Cecidomyiidae (Itonididae). Moth or Drain flies Moth flies, often called drain flies, are small, about 1/8 inch in length and often dark-colored. Their wings are covered with fine hairs, which give them a moth-like appearance. These flies rest on surfaces with their wings held over their back in a roof-like manner, and they have wing veins that extend in a parallel arrangement from the base to the tip of the wing, a pattern unique to the Psychodidae family of flies. They are weak flyers and exhibit a characteristic flying behavior of short hopping flights. The combination of characteristics described above can be used to distinguish moth flies from other flies in and around homes and other buildings.

Like all flies, moth flies undergo complete metamorphosis with egg, larval, pupal, and adult stages. Female moth flies lay eggs in moist to nearly saturated organic matter. In an urban environment, moth fly development often occurs in the slimy organic matter coating sink or shower drains, giving these flies an alternate common name “drain flies” used by many pest management professionals. However, moth flies may also be found developing in wet animal manure, sewage or even compost. Very large numbers of these flies in one area probably indicate a development site bigger than a few indoor drains. Once the eggs have been laid, they hatch in about 48 hours and continue to develop in the wet organic matter as larvae.

Moth fly larvae in the final (third) larval stage are approximately ¼ inch in length, have a distinct head, and a siphon on one end, which allows them to breathe in the wet environment. Immature flies pupate at their developmental site before emerging as adult flies. The life cycle of moth flies can be completed in as little as 8 days but can take as long as 24 days depending on temperature.

REFERENCES Aylsworth, Jean. 1993. Biological controls catch on with growers. Greenhouse Grower. December. pp. 77-78, 80-81. Chase, A.R. 1998. New bactericides and fungicides for disease control on ornamentals. Greenhouse Product News. December. pp. 22-24. Dr. Harry Hoitink Department of Plant Pathology Ohio Agricultural Research and Development Center The Ohio State University 1680 Madison Avenue Wooster, OH 44691-4096. Hoitink, Harry A., and Peter C. Fahy. 1986. Basis for the control of soilborne plant pathogens with composts. Annual Reviews of Phytopathology. Vol. 24. pp. 93-114. Dutky, Ethel. 1995. Here’s how to cut your losses due to disease. GMPro. October. p. 63-65. Gamliel, A. et al. No date. Solarization for the Recycling of Container Media. The Hebrew University of Jerusalem, Rehovot, Israel. Unpublished manuscript. 8 p. Garibaldi, Angelo, and M. Lodovica Bullino. 1991. Soil solarization in Southern European countries, with emphasis on soilborne disease control of protected crops. pp. 227-235. In: Jaacov Katan and James E. DeVay (ed.) Soil Solarization. CRC Press, Boca Raton, FL. Gentile, A.G., and D.T. Scanlon; Revised by Tina Smith. 1992. A Guide to Insects and Related Pests of Floricultural Crops in New England: For Commercial Growers. University of Massachusetts Cooperative Extension System. 36 p. Giblin, R.M., and S.D. Verkade. 1987. Solarization of small volumes of potting soil for disinfection of plant-parasitic nematodes. p. 174-176. In: Proc. Fla. State Hort. Soc. Vol. 100.

Gindrat, D. 1979. Biological soil disinfection. p. 253.287. In: D. Mulder (ed.) Soil Disinfection. Elsevier Scientific Publishing Co., New York, NY. Hoitink, H.A.J., Y. Inbar, and M.J. Boehm. 1991. Status of compost-amended potting mixes naturally suppressive to soil-borne diseases of floricultural crops. Plant Disease. September. pp. 869-873. Horiuchi, Seizo. 1991. Soil solarization in Japan. pp. 215, 218-223, 225. In: Jaacov Katan and James E. DeVay (ed.) Soil Solarization. CRC Press, Boca Raton, FL. Klassen, Parry. 1993. Mulling over methyl bromide. Greenhouse Grower. August. p. 118 & 120. Kuack, David. 1995. Janet Bandy on implementing an effective IPM program. Greenhouse Management and Production. April. pp. 56-57. Lindquist, Richard K. 1998. Evaluations of non-conventional pesticides for insect and mite control on greenhouse ornamental plants. Greenhouse Product News. July. pp. 52-55. Mahrer, Yitzhak. 1991. Physical properties of solar heating of soils by plastic mulching in the field and in glasshouses and simulation models. pp. 75, 81-86. In: Jaacov Katan James E. DeVay (ed.) Soil Solarization. CRC Press, Boca Raton, FL.

4: Integrated Pest Management and Insect Biology INTEGRATED PEST MANAGEMENT Consumer and grower concerns about widespread pesticide use, possible health risks from pesticide residues, problems with insecticide resistance, and groundwater contamination have led to increased interest in pest management programs that reduce use of broad spectrum, non-selective pesticides. A pest management program based upon Integrated Pest Management (IPM) strategies helps address these issues. Successful IPM programs combine accurate pest identification and scouting with cultural, biological and chemical controls in an economically and ecologically sound manner. Pest Identification Pest management decisions are initially based on correct/accurate identification and understanding of the arthropod (insect or mite) pest’s life cycle (egg to adult). Effective pest management depends on a greenhouse grower’s ability to determine which life stages are present and susceptible to pest management tactics. For example, spraying a pest control material (in this case an insecticide) to manage whiteflies is most effective when they are in the nymphal stages. Mis-identification of arthropod pests or their life stages can be costly and lead to inadequate control such that arthropod pest populations increase to levels that result in crop damage. Arthropod pest identification may be improved by participating in state-wide workshops and IPM training programs.

IPM SCOUTING AND DECISION-MAKING PRE-CROP SITE EVALUATION

One month before introducing a crop, evaluate the entire greenhouse and surrounding area. Before introducing a crop into a greenhouse it is imperative to remove weeds, algae, “pet plants,” and any plant and growing medium debris located throughout the greenhouse, particularly underneath benches, because these may provide refuge for certain arthropod pests. In addition, repair any drainage problems that may contribute to recurring arthropod pest outbreaks. A fallow period (with greenhouses empty of crops, and weeds) of at least four weeks may help to reduce pest pressure for the upcoming growing season. A break in production of even two weeks may be helpful in reducing pest pressure. Previous pest problems in the greenhouse and current management strategies should then be reviewed. A plan of action can be developed to avoid or reduce these pest problems. Prevention of arthropod pest problems may be easily accomplished if greenhouse growers take the time to identify, analyze and correct problems before crops are introduced. Scouting Scouting is the regular inspection of crops for insects, mites, diseases, and cultural problems. The individual responsible for scouting could be an employee or an outside consultant. However, for employee scouts, it is best that scouting be the acknowledged responsibility, so that routine greenhouse tasks do not interfere with any scouting duties. Scouting Tools Helpful scouting tools include a 10x to 20x hand lens, Optivisor™, dissecting microscope, digital camera, sticky cards, flagging tape (of different colors), scouting forms, pH and EC (electrical conductivity) meters, disease detection kits (see www.agdia.com), and resource information (listed at the end of this section). Inspection of Incoming Plants Inspect incoming plant material for the presence of insects, mites, diseases, or cultural problems such as nutritional deficiencies. If feasible, quarantine infested or problematic plants in an isolated greenhouse or area so they can be treated with a pest control material (insecticide or miticide) before they are placed in production areas.

Scouting Program Regular weekly scouting consists of using colored sticky cards, potato disks (to monitor for fungus gnat larvae), random plant inspections, plant tapping and use of indicator plants. Yellow and Blue Sticky Cards Yellow sticky cards are commonly used in greenhouses to scout for or monitor insect pest populations. These cards capture adult whiteflies, thrips, fungus gnats, shore flies, leafminers, and winged aphids. Remember that mites, mealybugs, scales, and nonwinged aphids don’t fly, so they are not captured on sticky cards. Also, it is important to note that many beneficial insects including parasitoids and predators may also be caught on yellow sticky cards. Position yellow sticky cards throughout the greenhouse, using approximately 3 to 4 cards per 1,000 ft2, or a minimum of one card per 1,000 ft2, with additional cards placed near openings such as doors, vents and sidewalls. Use clothespins and stakes to vertically attach sticky cards 4 to 6 inches (10 to 15 cm.) above the crop canopy. As plants increase in height, move the sticky card upward (vertically) on the stake. Blue sticky cards are more attractive to thrips (and even shore flies) and may be used to detect low thrips populations on susceptible crops such as impatiens and begonias. However, thrips and other insect pests captured on yellow sticky cards are easier to observe than on blue sticky cards. When monitoring for fungus gnat adults, place yellow sticky cards horizontally near the growing medium surface because more fungus gnat adults will be captured compared to placing sticky cards vertically above the crop canopy. Potato Disks Potato disks are used to monitor for fungus gnat larvae. Cut a fresh potato into disks 1.0 inch (2.5 cm) in diameter and ¼ to ½ inch (0.6 to 1.2 cm) in thickness; then press the disks into the growing medium surface in tagged or flagged pots. For plug trays, potatoes may be cut into small “French fry” shapes or wedges and inserted into the growing medium. In general, use 5 to10 potato wedges per 1,000 ft2 of greenhouse production area. After two days, inspect the undersides of the potato disks and/or wedges for the presence of fungus gnat larvae, which have distinct black head capsules.

Record the number of larvae located on each potato disk or wedge, and those present on the surface of the growing medium. Random Plant Inspections In order to detect problems early, it is important to inspect crops at least weekly for feeding damage and/or the presence of arthropod pest populations. When inspecting plants, select a set number or group of plants throughout the greenhouse (the actual number depends on the time and desired level of accuracy). Inspect leaves, stems and roots. Inspect plants at floor, bench and hanging basket levels. Sampling a pre-determined number of each type of plant increases the likelihood of locating “hot spots,” which are areas with high arthropod pest populations. Take advantage of previous experience by focusing on plant species that tend to be susceptible to arthropod pests. Inspect plants for both pests and natural enemies (if applicable). If inspecting plants during the day, keep in mind that certain insect pests (such as black vine weevils or Asiatic garden beetles) and natural enemies such as Atheta (=Dalotia) or Aphidoletes are more active at night. Plant Tapping Tap plant foliage or flowers over a sheet of white paper to monitor for arthropod pests such as aphid nymphs, thrips, mites, plant bugs, and leafhoppers. Indicator Plants Indicator plants are typically used to determine the effectiveness of pest management tactics or to monitor for the viruses (tospoviruses) such as impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV), which are vectored by the western flower thrips (Frankliniella occidentalis). Before implementing any pest management strategy, select and tag (or flag) the leaves or stems of 1 to 5 infested plants per 1,000 ft2. Afterward, inspect the indicator plants to assess if arthropod pests have been killed. This makes it possible to evaluate the effectiveness and longevity of control or regulation. To detect viruses transmitted by thrips, use either dwarf fava bean (Vicia faba) plants or ‘Summer Madness’ petunias. Position a blue card with the sticky portion covered near the indicator plants in order to attract adult thrips. If thrips adults possess any tospovirus, a brown, necrotic spotting will be

observed near white feeding scars within 48 hours. Rogue-out any infected petunia or fava bean plants, to remove any potential viral sources. Virus infections are systemic in fava bean but not petunia. Making Pest Management Decisions Each week, review scouting records to assess the effectiveness of your pest management tactics. Early detection of arthropod pests helps prevent the need to deal with extensive populations that may cause crop damage. It is also helpful to review scouting records at the end of each growing season to determine which arthropod pests were a problem and which pest management tactics provided the most effective control or regulation.

PEST MANAGEMENT IN GREENHOUSES Biological Control Biological control is the use of living organisms (natural enemies) such as insects, mites, fungi or bacteria, to control or regulate pest populations. Greenhouses provide suitable temperatures, relative humidity and light for numerous biological control agents or natural enemies including parasitoids (parasitic wasps), predators, and entomopathogenic nematodes. Greenhouses in the northern regions of the U.S. are typically not subject to massive migrations of insect pests from outdoor crops. Greenhouse environment conditions including temperature, light, and relative humidity during the winter are often stable, which may influence the performance and persistence of natural enemies. Many natural enemies are, in fact, commercially available and can be incorporated into existing greenhouse pest management programs. Some advantages of biological control agents include less worker exposure to toxic pest control material residues, less chance of spray damage, no re-entry intervals (REI), and lower risk of environmental pollution. Biological control is an important component of a resistance management program. Ornamental crop production is complex because of multiple arthropod pests, potential biological control agents, and pest management strategies. Natural enemies cannot be used in the same manner as pest control materials (insecticides or miticides). Pest control materials are typically applied after arthropod pests reach damaging levels. Effective pest control chemicals quickly reduce

arthropod pest populations. Using natural enemies to cure or regulate pest populations is less successful than applying them preventively. Natural enemies should be released early in the cropping cycle when plants are small, arthropod pest populations are low, and before crop damage occurs. Multiple releases of natural enemies may be required throughout the growing season in order to manage arthropod pests at low population levels. Guidelines for Success A biological control program will only succeed if these steps are followed: 1. correctly identify all arthropod pests; 2. purchase natural enemies from a reliable biological control supplier; 3. make sure there is a consistent supply of high quality natural enemies; 4. emphasize that proper shipping procedures be followed; 5. obtain directions from biological control suppliers on proper release rates and timing of applications; 6.

consult with suppliers of plant material to ensure that there are no longlasting pest control material residues on incoming plant material. In fact, request a list of pest control materials applied from plant suppliers;

7. establish a regular, consistent scouting program; 8. develop a strategy based upon greenhouse production plans; 9. obtain full commitment of the owner and/or manager; and 10.

ensure proper communication among all staff, employees and management regarding biological control programs.

Start any new biological control program in a small isolated greenhouse, in propagation houses, or in a greenhouse where edible crops such as herbs are being grown. This approach allows for gaining experience and then expanding into other production areas. It is critical to implement a scouting program and establish a favorable relationship with your biological control supplier early. The success of any biological control program relies on patience and a strong commitment to detail with an emphasis on scouting and record-keeping. Arthropod pest identification is extremely important when initiating

biological control programs in greenhouses because natural enemies, particularly parasitoids, are specific in the types of insect pests they use as hosts. For example, the aphid parasitoid Aphidius colemani attacks both the melon/cotton aphid (Aphis gossypii) and the green peach aphid (Myzus persicae), but does not attack the foxglove aphid (Aulacorthum solani). For arthropod pest identification information, consult trade journal articles, books, manuals, fact sheets, and picture identification guides, or send specimens to your Extension entomologist. Planning Timeline Plan carefully to ensure the success of your biological control program. If you haven’t used biological controls before, start planning 6 months to one year in advance. Contact suppliers; review your current pesticide usage and move toward using pesticides that have a shorter residual for this transition period. Contact your plant suppliers before receiving plant materials,to determine what pesticides have been used on cuttings or liners. Schedule delivery of the natural enemies, and consult with your supplier to decide whether a regular standing order or week-by-week order is needed. Product Quality Natural enemies are living organisms that must be handled and stored carefully in order to maximize survival and sustain viability. Two important considerations associated with natural enemy quality are shipment arrival time and survivability upon arrival. Shipments of natural enemies should be received within four days after placing an order. The package containing the natural enemies must be shipped in a sturdy container such as a polystyrene box that minimizes exposure to high and low temperatures. Request that the biological control supplier include ice packs and a data logger (if possible). Also, make sure the container is secured with good packing material during shipment. If appropriate, ask your biological control supplier(s) how to best evaluate incoming shipments. Biological control suppliers often send a description of what to look for when one receives the natural enemies. Upon receipt of natural enemies, check the quality of the shipment. Assess the number of shipment days and how cold or warm the ice packs are. When you open the shipping container and retrieve the packages of natural enemies, consider the following: 1) Did you receive the correct natural enemy? That is, did you acquire what you ordered? 2) Did you receive the correct number of

packages or units? Is each package or unit labeled with the estimated number of natural enemies? 3) Are the natural enemies alive? If a majority (>50%) of the natural enemies are dead, contact the biological control supplier immediately and request another shipment. Be sure to return the original shipment to the supplier. If natural enemies shipped are mobile (for example, predaceous mites and some parasitoids), inspect them to determine if they are actively moving. To check natural enemies that are shipped as eggs or pupae, place a small sample in a shaded, unsprayed area for 2-3 days and then look for the active larvae or adults. Below are procedures to assess the quality of predatory mites, whitefly parasitoids, and entomopathogenic nematodes. Predatory Mites After receiving a shipment of predatory mites such as Phytoseiulus persimilis or Neoseiulus (Amblyseius) cucumeris, evaluate a small sample of the contents to see whether the mites are alive, and whether you received the correct quantity. For example, if by volume, the entire package fills five Styrofoam coffee cups, select 1/8 cup or 2.5% of the material, after first gently rotating the contents of the package to mix the predatory mites with the bran carrier. Pour the 1/8-cup of material onto a sheet of white paper (8 × 11 inch or 20 × 28 cm) and gently spread the contents out using a soft camelhair paint brush. Count the number of mites that move to determine the viability of the package. If no active mites are detected, immediately contact your biological control supplier to discuss appropriate procedures. Shipments of N. cucumeris may also contain grain mites (Acarus siro), which serve as a food source. Grain mites are white, have long protruding hairs on their backs and are generally less mobile than the tancolored N. cucumeris. Avoid counting any grain mites during the quality evaluation process. You should also check the rim of the container in which the predatory mites are shipped for the presence of live individuals. Whitefly Parasitoids Shipments of whitefly parasitoids contain either paper cards with a predetermined number of pupae per card (e.g., 100 pupae per card for Encarsia formosa) or an estimated number of pupae in loose bran or sawdust (e.g., Eretmocerus eremicus). To assess whitefly parasitoids shipped as pupae

on paper cards, place a single card into a glass jar with a tight-sealing lid. Expose the jar to room temperature (70 to 75ºF or 21 to 24ºC) and position out of direct sunlight. Do not disturb the jar for approximately 5 days. After this time, count or estimate the number of adult parasitoids that have emerged from the pupae. Make sure e”95% of adult parasitoids have emerged. For E. eremicus, which is sold as loose pupae, place a subsample (approximately 2% by volume of the total contents) into a jar with a tight-sealing lid. After storing at the same temperatures and time as described above for E. formosa, count the number of live adult parasitoids that emerge from the pupae. Again, make sure e”95% of adult parasitoids emerge. If 90%. This fungus may not be compatible with the convergent ladybird beetle (Hippodamia convergens), depending on the concentration of spores applied.

B.

Isaria (=Paecilomyces) fumosoroseus: The entomopathogenic fungus, Isaria (=Paecilomyces) fumosoroseus is also commercially available under the trade names NoFly WP and Preferal. This fungus is most effective when the relative humidity is 80% or higher for 8 to 10 hours.

Pest Control Materials Pest control materials (insecticides) with contact, translaminar, or systemic activity can be used to control or regulate aphid populations. Translaminar means that after application the material penetrates leaf tissues and forms a reservoir of active ingredient within the leaf. This provides extended residual activity even after spray residues dissipate. It is important to rotate insecticides with different modes of action to delay the onset of resistance. Insecticide applications must be initiated early in the cropping cycle, when plants are small. The nymphs of some aphid species reside between the scales of leaf buds or in flowers. This reduces their exposure to contact insecticides, making repeat applications necessary. A surfactant may improve coverage of plant parts when used with either wettable or soluble powder formulations. In some instances, insecticidal soaps and/or highly

refined horticultural oils may provide control of aphids, particularly when populations are low. However, since these insecticides kill exclusively by contact and have minimal residual activity, thorough coverage of all plant parts is essential. Insect growth regulators and pyrethroid-based insecticides may also provide control or regulation of aphids. Systemic insecticides, including the neonicotinoidbased insecticides imidacloprid (Marathon® and many generic products), thiamethoxam (Flagship®), acetamiprid (TriStar®), and dinotefuran (Safari®), effectively control or regulate populations of aphids for extended periods of time when applied early in the cropping cycle. The same is true of the selectivefeeding blockers, pymetrozine (Endeavor®) and flonicamid (Aria®). A number of insecticides have both translaminar and systemic properties. Beetles Identification, Biology and Life Cycle Beetles are a large group of insects characterized by hardened forewings. Both adults and larvae have chewing mouthparts that cause damage to a wide range of plants. Leaf feeding beetles such as the lily leaf beetle and scarab beetles are important pests in herbaceous perennial production. Other damaging beetles include tortoise beetles, flea beetles, various spotted and striped cucumber beetles, larvae of click beetles (wireworms), and blister beetles. During weekly plant inspections, look for chewed leaves, or pinholes from flea beetle feeding. Scarab Beetles Scarab beetles are large brightly colored beetles with lamellated tips on their antennae. Asiatic garden beetles, Oriental beetles, and Japanese beetles feed on many species of herbaceous perennials, woody ornamentals and vegetables. However, European chafer adults are not foliage feeders. The fleshy legless larvae, which are known as “white grubs,” develop on the roots of many plant species. Identification of the particular grub species is important because the effectiveness of pest control materials and beneficial nematodes against white grubs vary according to species. Larvae can be identified by the pattern of hairs (“rasters”) on the tip of their hind end. A.

Asiatic garden beetle (Maladera castanea) adults are about 3/8 inch (0.95 cm)long and are cinnamonbrown in color. They

are often found near the rootsof plants. Asiatic garden beetles feed at night on aquilegia, aster,chrysanthemum, dahlia, delphinium, helianthus, heuchera, phlox,physostegia, rose, rudbeckia, salvia and zinnia. Their nighttime feedingcauses Cshaped notches on the edges of leaves. During the day, adultsburrow into mulch or soil/growing medium, or may be found under pots.“White grubs” feed on the roots of grasses and flowering plants. Asiaticgarden beetles overwinter as grubs in the soil/growing medium and adultsemerge the following summer (mid-July to mid-August). There is onegeneration a year. Contact insecticides may be applied against the adults.Repeat applications may be needed. B.

Oriental beetle (Anomala orientalis) adults are about 1/2 inch (1.27 cm) long,dark brown or strawcolored, and have dark markings on their wing covers.Adults emerge from the soil/growing medium in mid- June and are presentuntil August. They are active during the day and night. Adults do very littlefeeding on plant leaves. The “white grubs” feed on the roots of herbaceousperennials and woody ornamentals.

C.

Japanese beetle (Popillia japonica) adults are 1/3 to ½ inch (8.4 to 12.6 mm)long, metallic green with coppery wing covers and white tufts of hair nearthe end of the abdomen. Adults feed during the day on many woody andherbaceous ornamental plants. Adults emerge from the soil/growingmedium in June and July and feed for about 30 to 45 days. Eggs are laid inthe soil in grassy areas and hatch into white, C-shaped grubs that feed onturf grass roots. Japanese beetles overwinter as grubs in the soil/growingmix below the frost line. There is one generation per year.

Scouting Japanese beetles are extremely mobile, and once feeding begins, they emit feeding or aggregation pheromones to attract other beetles to their location. Look for feeding between leaf veins (called “skeletonization”) on favored hosts. Also, check for “white grubs” in the soil or growing medium. Cultural Control

Weed control in and around production areas helps to eliminate potential alternative food sources. Shade cloth can be used to exclude adults from hoop houses. Japanese beetles are strong fliers. Japanese beetle traps attract adult beetles, but are not recommended because they may increase feeding damage. Biological Control The female winsome fly (Isocheta aldrichii) is a natural parasitoid of adult Japanese beetles. Look for distinct white eggs on the thorax of adult beetles. The spring Tiphia (Tiphia vernalis) is a parasitoid that attacks Japanese beetle and Oriental beetle grubs. A recent survey in Connecticut found that spring Tiphia wasps are widely distributed. Beneficial nematodes (Heterorhabditis sp.) are commercially available for use against “white grubs.” Scarab beetle species vary in their susceptibility to infection and “white grubs” may defend themselves against nematode infection. Pest Control Materials Apply contact insecticides as soon as adult beetles are observed. However, many contact insecticides may be harmful to bees, predatory mites and insects. To control “white grubs”, apply insecticides in grassy areas surrounding production areas.

ADDITIONAL LEAF-FEEDING BEETLES Lily Leaf Beetle The lily leaf beetle (Liloceris lilii) was introduced into the U.S. in 1992, and has since spread throughout New England. Adults are ¼ to 3/8 inches (6.3 to 9.5 mm) long and bright scarlet-red in color, with black legs, head and antennae. Larvae are orange, brown, or yellow. They resemble a fragment of soil, as they transport their excrement on their backs. Both adults and larvae feed on species of Fritillaria, Lilium, Polygonatum and Nicotiana. Overwintering adult beetles emerge from the soil or growing medium in early spring. Females lay up to 250 eggs over two growing seasons, on the underside of lily leaves. Larvae, which feed for approximately two weeks before entering the soil to pupate, cause most of the damage to plants. Adults emerge from pupae in 3 to 4 weeks and feed on plants until fall. Adults overwinter in soil/growing medium and plant debris.

Tortoise Beetle Golden tortoise beetle (Metriona bicolor) adults are shiny, golden beetles, less than ¼ inch (6.3 mm) long with thin margins that extend out from their body and a shield-like structure covering the head. They are sometimes mistaken for ladybird beetles and are also known as “goldbugs.” Tortoise beetles overwinter as adults and there is one generation per year. During late spring and early summer, adults lay their eggs on leaves. Eggs hatch into yellow to brown, oval, broad, spiny, flattened larvae that use their rear spines to hold debris and excrement over their back. Both adults and larvae feed on plants in the morning glory family (Convolvulaceae) including the sweet potato vine (Ipomoea spp.), causing distinct round feeding holes. Contact insecticides labeled for leaf-feeding beetles may be applied in production areas where damage is unsightly. Caterpillars Identification, Biology and Life Cycle Caterpillars are the larval stage of butterflies and moths. Some caterpillars are large (1 to 2 inches or 2.5 to 5.1 cm in length) whereas others, particularly when young (1st or 2nd instar), may not be visible without the aid of a 10X hand lens. Caterpillars have five or fewer pairs of prolegs (seen on the abdomen) and hooks called crochets at the tip of their prolegs. Caterpillars that may be encountered in greenhouses include armyworms, cutworms, imported cabbage worm (Artogeia rapae), diamondback moth (Plutella xylostella), leaftiers, leafrollers (Choristoneura spp.), loopers, tobacco budworm (Helicoverpa virescens), salt marsh caterpillar (Estigmene acrea), and European corn borer (Ostrinia nubilalis). Damage is only caused by the larvae, which feed on leaves, stems, and flowers whereas adults feed on nectar or pollen. Infestations typically begin when infested plants are introduced into a greenhouse. In addition, because adult moths such as cabbage looper (Trichoplusia ni) are active at night and are attracted to lights, females may enter greenhouses and lay eggs on plants. Adult European corn borer females sometimes migrate from nearby cornfields and lay eggs on garden chrysanthemums and herbaceous perennials in outdoor production. The emerging larvae feed on these crops. European corn borer larvae are ¾ to 1.0

inch (2.0 to 2.5 cm) long, and are cream colored with brown spots. Mature larvae overwinter in the stems of host plants. In New England, there are one to two generations per year. University vegetable specialists use pheromone traps to monitor European corn borer flight activity in Connecticut and Massachusetts. Consult online vegetable pest messages and newsletters for this information, which may be helpful in timing insecticide applications. The European pepper moth (EPM) (Duponchelia fovealis) is a relatively new invasive species, not yet detected in New England. It has been reported on poinsettia, geranium and begonia. Look for the larvae, girdling of stems, and rolled leaves near the base of plants. Scouting Scouting for caterpillars is important and avoids having to deal with large populations that may damage crops. Inspect plants routinely for signs of feeding damage and the presence of fecal pellets (caterpillar frass). Begin plant inspections when adults are flying. When scouting, check plants closest to greenhouse openings where adults may enter, especially areas closest to vegetable fields. Cultural Control Eliminate weeds that may serve as alternative hosts. Cleaning up plant debris may help remove overwintering pupae. Install insect screening over openings such as vents and sidewalls to prevent the entry of adults into the greenhouse. Microbial and Biological Control A.

Bacillus thuringiensis subsp. kurstaki: Spray applications of the soil-bornebacterium Bacillus thuringiensis subsp. kurstaki, also known as Btk and soldcommercially as Dipel Pro DF®, Deliver®, and Javelin®, kill youngcaterpillars. This bacterium must be consumed by the caterpillar in orderto be effective, and thorough coverage of all plant parts is essential. Becausethe bacterium is susceptible to ultraviolet light degradation, repeatapplications are usually necessary. Caterpillars typically stop feeding within24 to 48 hours after eating the bacterium, and die after 3 to 4 days.

B.

Trichogramma: Parasitoids in the genus Trichogramma only attack the eggstage of various caterpillar species including diamondback moth, cabbagelooper and imported cabbageworms. They do not parasitize the larval orcaterpillar stage.

Fungus Gnats Identification, Biology and Life Cycle Fungus gnat (Bradysia spp.) populations commonly develop in moist environments such as propagation greenhouses. The larvae are translucent to white in color, legless, and approximately ¼ inch (6.3 mm) long when mature. They have a distinct black head capsule. Adults resemble mosquitoes, and are 1/8 inch (3.1 mm) long, with long legs and antennae, and their forewings display a “Y”-shaped vein pattern. Fungus gnat adults are weak fliers and may be observed resting on the growing medium surface or moving across leaves in the lower plant canopy. Adults are primarily a nuisance when present in large numbers; however, they have been implicated in carrying Botrytis spores on their bodies. Larvae feed on fungi and decaying organic matter but they also feed on plant roots, reducing the plants’ ability to take up water and nutrients. The larvae also tunnel into the crowns and stems of plants. This burrowing activity creates wounds that allow soilborne pathogens to enter, and can kill plants. Fungus gnat larvae may also vector soil-borne pathogens such as Pythium, Thielaviopsis and Fusarium. Fungus gnats are a common problem on geraniums, begonias, poinsettias, sedums and bulb crops, especially if the growing medium contains a high percentage of composted bark or peat moss. Female fungus gnats can lay up to 200 white eggs in clusters of 20 to 30 on the surface or in the crevices of moist growing media, particularly those with high organic matter content. Eggs hatch in 5 to 6 days. The larvae feed on plant roots for approximately 14 days before transitioning into pupae. Fungus gnats remain in the pupal stage for 5 to 6 days before adults emerge. Adults live approximately 10 days. The life cycle from egg to adult typically takes 21 to 28 days, depending on temperature. The presence of overlapping generations can make control or regulation difficult. Scouting

Monitor for fungus gnat adults by placing yellow sticky cards at the base of plants, either on the growing medium surface or on the edge of flats. Inspect yellow sticky cards weekly to detect early fungus gnat infestations. Keep detailed records of the numbers of adults captured on yellow sticky cards to determine the efficacy of pest management tactics. Monitor for fungus gnat larvae by inserting potato disks or wedges into the growing medium (discussed previously). Ten potato disks may be sufficient to monitor a 10,000 ft2 greenhouse. Check disks after 48 hours, and count the number of larvae on each disk and any that are present on the growing medium surface. Replace old disks with new ones. Inspect young cuttings for signs of fungus gnat feeding and inspect root systems for fungus gnat larvae. Cultural Control Good sanitation practices that reduce breeding sites can prevent or minimize problems with fungus gnat populations. Avoid overwatering and allowing moisture to accumulate underneath benches. Be sure to solve drainage problems. Remove old growing medium and plant debris from inside and around the greenhouse. Inspect incoming plugs for fungus gnat larvae, if feasible, and fungus gnat adults. Fungus gnats may enter greenhouses in commercial bagged growing medium or on infested plants. Covering the growing medium with a layer of either coarse sand or diatomaceous earth does not inhibit adult emergence or prevent egg-laying by adult females. Diatomaceous earth absorbs moisture from the growing medium, resulting in the development of cracks that provide sites where larvae can pupate and females can lay eggs. Biological Control Several commercially available natural enemies control or regulate fungus gnat larval populations, including the soil-dwelling predatory mite, Hypoaspis miles (=Stratiolaelaps scimitus); the entomopathogenic nematode, Steinernema feltiae; and the rove beetle, Atheta coriaria (=Dalotia coriaria). All three natural enemies are most effective if applied before fungus gnat populations are large. In addition, the growing medium should be thoroughly moist before applying any of these natural enemies. Be sure to consult biological control suppliers for information pertaining to release rates. Below are descriptions of each of these natural enemies: Predatory Mites

Hypoaspis miles (=Stratiolaelaps scimitus): This soil-dwelling generalist predatory mite not only feeds on fungus gnat larvae but may also feed on thrips pupae and shore fly larvae. This mite prefers to feed on first instar fungus gnat larvae. If prey is not available then H. miles feeds on plant debris and algae. It is important to initiate releases early in the growing season before fungus gnat larval populations are high. Applications can also be directed to the soil under greenhouse benches. Avoid mixing H. miles (=Stratiolaelaps scimitus) into growing media prior to planting because this decreases survival. Applications must be initiated after planting and the growing medium should be moist but not saturated. H. miles (=Stratiolaelaps scimitus) is active when growing medium temperatures are e”50ºF (10ºC). In order to evaluate this mite’s effectiveness, look for reductions in numbers of fungus gnat adults on yellow sticky cards and larvae on potato disks. Pest control materials (insecticides, miticides and fungicides), when applied as wet sprays to foliage with high-volume spray equipment, may enter the growing medium and directly or indirectly affect H. miles. For example, H. miles (Stratiolaelaps scimitus) is negatively affected when exposed to chlorpyrifos (DuraGuard®), but pyriproxyfen (Distance®), novaluron (Pedestal®), fosetyl-aluminum (Aliette®) and mefenoxam (Subdue®) do not directly harm this predatory mite. Predatory Beetles Atheta coriaria (=Dalotia coriaria): This generalist predator feeds on fungus gnat and shore fly larvae, and supposedly thrips pupae, in the growing medium. Rove beetle adults are slender, dark brown to black, and covered with hairs. The adults are 1/8 inch (3.1 mm) long with very short wing covers. They fly to disperse throughout a greenhouse from original release sites. Larvae are cream to brown in color, depending on age. Both stages inhabit cracks and crevices in the growing medium. Adults and larvae prey on fungus gnat eggs and larvae. Once established in a greenhouse, rove beetles may be present yearround, although populations may fluctuate depending on fungus gnat larval populations. Because rove beetles are generalist feeders, they may consume other natural enemies such as H. miles (=Stratiolaelaps scimitus). In addition, soil-dwelling predatory mites may feed on young rove beetle

larvae. Rove beetles are commercially available as adults from most biological control suppliers. Consult your supplier about release rates. Temperatures of 65 to 80ºF (18 to 26ºC) and a relative humidity of 50 to 85% are optimal for survival. Adults are nocturnal, so they are best released in the evening. Both adults and larvae are difficult to detect by scouting since they tend to hide in the cracks and crevices of growing medium. Adults can be observed on the surface of the growing medium with their abdomens raised. The pest control materials dinotefuran (Safari®) and thiamethoxam (Flagship®) are directly toxic to rove beetle adults, whereas the insecticides Bacillus thuringiensis subsp. israelensis (Gnatrol WDG®), azadirachtin (Ornazin® and other generic products), diflubenzuron (Adept®), and the fungicides azoxystrobin (Heritage®), fosetyl-aluminum (Aliette®) and metalaxyl (Subdue®) are compatible with rove beetle adults. Rove beetles are also compatible with entomopathogenic nematodes. Entomopathogenic Nematodes Steinernema feltiae: This entomopathogenic or beneficial nematode attacks fungus gnat larvae. Infective juveniles enter the host body through natural openings such as the mouth, anus and breathing pores (spiracles) and then release a symbiotic bacterium (Xenorhabdus spp.) that kills fungus gnat larvae by dissolving their internal contents. Infected fungus gnat larvae may be opaquewhite to light yellow. Larvae are killed within 1 to 2 days. After the nematodes reproduce and multiply in the dead cadaver, the infective juveniles exit and search for new hosts. Nematodes are typically applied as a drench. They can also be applied through drip irrigation systems, but filters must be removed. Application Tips To assess the viability of shipments prior to application, place a small quantity of the product (5 mL) in a shallow container or Petri dish and add 1 to 2 drops of tepid water. After a few minutes, look for active infective juveniles, which have a slight “S” or “J” curvature at the end of their bodies. Follow label directions to determine the appropriate number of entomopathogenic nematodes to apply. Nematodes can also be applied through a fertilizer injector. It is essential to agitate the mixture in the stock solution with a small submersible pump so

that the nematodes do not settle to the bottom of the container. Remove the filters and use a large-holed watering nozzle like a water-breaker head. Treat crops with entomopathogenic nematodes early in the production cycle. Either apply directly to cuttings prior to sticking, or just after planting. Repeat applications may be required if fungus gnat populations are high. Growing medium temperatures must be 50 to 80ºF (10 to 26ºC), with optimum temperatures of 60 to 70ºF (15 to 21ºC). Irrigate the growing medium before and after applying nematodes. The nematodes require moisture in order to move within the pores of the growing medium. Apply the nematodes in the evening, at dusk or on cloudy, overcast days because the nematodes are extremely sensitive to desiccation in ultraviolet light. In general, entomopathogenic nematodes or beneficial nematodes are compatible with most pest control materials although those in the carbamate and organophosphate chemical classes may be both directly and indirectly harmful to entomopathogenic nematodes. Hunter Flies: Yellow sticky cards may capture hunter fly (Coenosia attenuata) adults, which either fly into unsprayed greenhouses during the growing season or are introduced on new plant material. Hunter fly adults resemble common house fly (Musca domestica) adults in appearance. They range in length from 1/8 to ¼ inch (0.31 to 0.63 cm). Males have pale yellow legs and females have black legs. Hunter flies may be misidentified as shore flies, but hunter flies have wings that are uniformly clear and may appear iridescent when they are observed in full sun perching on plant leaves. Hunter flies are also nearly twice as large as shore flies. In addition to fungus gnat adults, hunter fly adults attack and feed on shore fly, whitefly, and leafminer adults. Adult hunter flies only attack prey species that are flying. After adults capture prey in flight, they puncture the prey with their dagger-like mouthparts, and consume their prey’s internal body fluids. The soil-dwelling larvae are also predaceous and feed on fungus gnat larvae and other insects that reside in the growing medium. Hunter flies establish in greenhouses by inhabiting the growing medium. The parasitoid Synacra pauperi may also be captured on yellow sticky cards, especially in unsprayed greenhouses. Females insert eggs into fungus gnat larvae. Eggs hatch into larvae that consume the internal contents of fungus gnat larvae. Parasitized fungus gnat larvae eventually pupate, and then an adult S. pauperi emerges. The population’s maximum rate of increase is

higher than fungus gnat larvae at 73ºF (23ºC). Pest Control Materials Insect growth regulators, microbials and other pest control materials applied to the growing medium may be effective in controlling or regulating fungus gnat larva populations. Most pest control materials do not affect eggs or pupae, so repeat applications are typically required. The soil-borne bacterium Bacillus thuringiensis subsp. israelensis (Gnatrol WDG®) may be used before fungus gnat larva populations are high since the bacterium must be ingested in order to be effective. Applications are more effective on the young larvae (1st instar) than mature larvae (3rd and 4th instars). This material should be applied weekly until fungus gnat populations start to decline. Do not use this product if fungus gnat populations are excessive. In this case, use insect growth regulators such as pyriproxyfen (Distance®), cyromazine (Citation®), or diflubenzuron (Adept®). Leafhoppers Identification, Biology and Life Cycle

Fig. 4.2: Aster leafhopper. A - adult, B - front view of head, C - nymph

Leafhoppers are not common insect pests of greenhouses, although the potato leafhopper (Empoasca fabae) and the aster leafhopper (Macrosteles quadrilineatus), both pests of outdoor crops including cut flowers, woody ornamentals and herbaceous perennials, may be detected on yellow sticky cards inside greenhouses. Both leafhopper species are approximately 1/8 inch (3.0 mm) long, with slender wedge-shaped bodies. Adults hold their wings rooflike over their bodies. Nymphs resemble adults but lack wings. Leafhoppers are very active, particularly when disturbed. The adults and nymphs may walk (sideways, backwards or forward), jump or fly.

Leafhoppers possess piercing-sucking mouthparts which they use to withdraw plant fluids, causing stunting, leaf yellowing, distortion and loss of vigor. The aster leafhopper transmits the phytoplasma that causes aster yellows. Potato leafhopper females insert eggs into the midrib or larger veins of plant leaves, or into leaf petioles or stems of astilbe, dahlia, gaura, hollyhock and lupine. Eggs hatch in 10 days. Nymphs feed on plants and undergo five molts before transitioning into adults. Adults can live up to two months. The life cycle from egg to adult may be completed in approximately four weeks. Aster leafhopper is widespread throughout the eastern United States, and is carried northward each year on wind currents. Aster leafhopper overwinters as an egg, and in southern Ontario there can be up to five generations during each growing season. Sage leafhopper (Eupteryx melissae) and mint leafhopper (Eupteryx decemnotata) have also been observed on various herbs such as rosemary, sage, catnip, mint, lavender and oregano. They are small, brightly colored and distinctly marked leafhoppers. Scouting Look for the fast moving adults and nymphs on the underside of leaves. Their feeding causes stippling of foliage that resembles spider mite feeding, and stunting and distortion of new growth. Potato leafhopper also injects a toxin as it feeds, so that leaves develop a V-shape, brown edge burn at the tip is known as “hopperburn” that may be mistaken for leaf scorch due to drought stress. Their white skins may be present on the foliage after shedding. Yellow sticky cards are helpful to trap the fast moving adults, and make it easier to determine the actual species. Biological Control S limited number of natural enemies are commercially available for managing the fast moving leafhoppers. In some cases, lacewings may help suppress leafhopper populations. Pest Control Materials Control of leafhoppers with contact insecticides is difficult because they are very mobile, and new leafhoppers can enter treated areas after sprays have dried. Systemic insecticides may be applied to ornamental plants to prevent feeding damage when leafhoppers first appear.

Leafminers Identification, Biology and Life Cycle Leafminer species in the family Agromyzidae are common insect pests of greenhouse-grown crops. The species typically encountered in greenhouses are American serpentine leafminer (Liriomyza trifolii), serpentine leafminer (Liriomyza brassicae) and pea leafminer (Liriomyza huidobrensis). Chrysanthemum leafminer (Chromatomyia syngenesiae) is not common but is an important insect pest. Adult leafminers resemble houseflies and are ¼ inch (6.0 mm) long with black bodies and yellow heads. In addition, their bodies possess yellow markings that are useful to distinguish leafminer adults from shore fly adults. Adults may be observed walking on plant leaves and flowers. Females use their ovipositor (egglaying structure) to pierce young leaves, and then they feed on the liquid that exudes from the wounded leaf tissue. Females insert eggs into leaves. Larvae that emerge from eggs are initially 1/16 inch (1.5 mm) in length and 1/13 and 1/7 inch (2.0 to 3.5 mm) long when mature. They are typically bright yellow to brown. As larvae feed within the leaf tissues, they create either serpentine or blotched mines, depending on the species. In general, leafminer females lay approximately 100 eggs during their 2 to 3 week lifespan. Eggs hatch in 5 to 6 days, and larvae feed beneath the leaf cuticle for about two weeks. The final larval instar creates an opening in the leaf and then falls to the ground. Larvae burrow into the growing medium or soil underneath benches to pupate. Leafminers require complete darkness in order to successfully pupate. Pupae are brown in color, and adults emerge within a two-week period. The duration of the life cycle depends on the host plant and temperature. For example, it takes 64 days to complete the life cycle at 59ºF (15ºC) but only 14 days at 95ºF (35ºC). There can be multiple generations each growing season. Scouting Inspect incoming plants for signs of leafminer activity. When visually inspecting plants, it is important to look for any egg-laying punctures, which appear as white spots on the tops of leaves. Use yellow sticky cards to monitor leafminer adult populations, especially on susceptible plant material. Cultural Control

Avoid overfertilizing plants, particularly with nitrogen. Overfertilized plants are more attractive to adult females for egg-laying. Biological Control Commercial natural enemies for leafminers include two parasitoids, Diglyphus isaea and Dacnusa sibirica. Dacnusa may also be sold in combination with Diglyphus. Diglyphus isaea is a small (1/25 to 1/11 inch or 1.0– 2.3 mm long) ectoparasitoid (females paralyze the host or prey before laying one or more eggs adjacent to leafminer larvae). Adult females attack second instar larvae within leaf mines. Dacnusa siberia is 1/11 to 1/9 inch (2.5 to 3.0 mm) long with extended antennae. It is an endoparasitoid (females lay eggs directly into leafminer larvae) that uses odors emitted from leafminer frass to locate larvae within damaged plant tissue. Dacnus is better adapted to lower temperatures (60° F or 15° C) than Diglyphus. Both of these parasitoids are most effective in controlling or regulating leafminer populations in long-term crops such as cut-flowers and stock plants, and they should be used preventively to control or regulate low leafminer populations. When scouting, look for short mines with the dead leafminer larvae inside the mines. Avoid applying broad spectrum pest control materials in the organophosphate, carbamate and pyrethroid chemical classes since many of these materials leave residues that may persist and kill both parasitoids for several weeks. Release the parasitic wasps in the morning or evening. Remove yellow sticky cards before making releases, as parasitoids are attracted to them. Replace the cards 3–4 days after applying the parasitoids. Diglyphus isaea has proven to be effective in controlling or regulating leafminers in greenhouse-grown chrysanthemums and Transvaal daisy (Gerbera jamesonii). Pest Control Materials The use of insecticides to control or regulate leafminer populations may be difficult because several species have developed resistance to a number of commonly used insecticides. Applications of pyrethroid-based insecticides may be required every 3– 4 days to kill adults as they emerge from pupae in

the growing medium or soil. Apply in the morning, when when females are actively laying eggs, as wet sprays may disrupt their ability to deposit eggs into leaf tissue. Several pyrethroid-based insecticides have repellent properties that may deter adult females from laying eggs, thus minimizing damage to plant leaves. Insecticides with translaminar properties, including several insect growth regulators and neonicotinoids, are effective in killing larvae within leaf mines. Mealybugs Identification, Biology and Life Cycle Mealybugs are typically a problem on long-term crops such as cut flowers, orchids, and foliage plants. Species encountered in greenhouses include the citrus mealybug (Planococcus citri), longtailed mealybug (Pseudococcus longispinus) and obscure mealybug (Pseudococcus viburni). Longtailed mealybug can be a major problem on conservatory plants such as cycads, orchids, palms and ferns. It is easy to identify due to the presence of long waxy filaments that protrude from the end of its abdomen. Obscure mealybug also has waxy filaments but they are much shorter than those of longtailed mealybug. Citrus mealybug lacks any waxy filaments on the body and has a gray stripe that extends the length of the body. Mealybugs that have been introduced into the U.S. and which may also be present in greenhouses include the pink hibiscus mealybug (Maconellicoccus hirsutus), madeira mealybug (Phenacoccus madeirensis), and the Mexican mealybug (Phenacoccus gossypii). Mealybugs usually enter a greenhouse on infested plant material. Mealybugs use their piercing-sucking mouthparts to withdraw plant fluids. Both nymphs (referred to as crawlers) and adults feed on plants and cause stunting, leaf yellowing and distortion of plant parts. While feeding, mealybugs may inject toxic saliva into plant tissues, and excrete a copious amount of honeydew (clear, sticky liquid) that serves as an excellent growing medium for black sooty mold fungi. The presence of black sooty mold fungi inhibits plants’ ability to manufacture food via photosynthesis, and detracts from the plants’ aesthetic appearance. Mealybugs are soft-bodied, segmented, oval-shaped insects. Adult females are 1.0 to 5.0 mm long and are usually covered with a white, powdery, waxy material. A fringe of waxy filaments may be present on the

margin of their body.

Fig. 4.3: Adult mealybug

Mealybugs retain their legs throughout development,allowing them to move from plant to plant within agreenhouse. Newly emerged nymphs or crawlers maturein 6 to 9 weeks. Some mealybug species, like thelongtailed mealybug, give birth to live offspring. Otherspecies, such as citrus and obscure mealybug, lay massesof up to 100 yellow-to-orange eggs in a white cottony sac.Eggs hatch in 5 to10 days, but unhatched eggs or youngnymphs may remain inside the cottony sac if environmental conditions such as temperature and relative humidity are not favorable. Adult males of most mealybug species are small winged insects that lack functional mouthparts. Their primary role is to fertilize females. Scouting Early detection and isolation of infested plants is important to avoid mealybug outbreaks. Routine visual inspections of susceptible plants make it easier to deal with mealybugs with either pest control materials or natural enemies. Mealybugs feed on a wide range of plant hosts including coleus, croton, dracaena, English ivy, fuchsia, gardenia, hibiscus, mandevilla, stephanotis, palms, orchids, cacti and succulents. Mealybugs are usually located on leaf undersides, petiole and leaf junctions, and near the base of plants. They may also be present on the inside of container lips and in container drainage holes. Root-feeding mealybugs (Rhizoecus spp.) appear as masses of wax, and may occasionally be detected on the roots of wilting plants. These mealybugs, which are not as active as mealybugs that feed above ground, are typically covered with a fine, powdery, wax-like material. Root mealybugs do not possess the marginal filaments that are typical of other mealybugs. Nymphs are covered by a white waxy material, and are located in the crevices of growing media or in excavated chambers on the outer edge of the root ball.

Root mealybug feeding causes stunting and leaf yellowing. Damage is usually noticeable when high populations are present. Once introduced into a greenhouse, root mealybugs may spread to other plants in water that drains from containers, in growing medium or plant debris, or on equipment. Root mealybugs may also infest adjacent plants by crawling through drainage holes of containers. If a greenhouse is infested with root mealybugs, all infested plants and debris should be discarded immediately. Disinfect/disinfest contaminated containers before reusing them. Cultural Control Inspect incoming plants for signs of mealybugs. Start with clean plant material. Do not carry over stock plants or “pet plants” that may be mealybug-infested. Immediately dispose of severely infested plants. Biological Control Parasitoids may suppress or regulate mealybugs in conservatories and where plants are maintained for extended periods. Identify mealybugs to species before releasing any natural enemies. Anagyrus pseudocci, a parasitoid, attacks the citrus mealybug, parasitizing both the second and third larval stages and adults. In general, mealybug predators are less efficacious than mealybug parasitoids. The predatory ladybird beetle Cryptolaemus montrouzieri, also called the “mealybug destroyer,” may be used to control or regulate citrus mealybug populations. Both adults and larvae are predaceous upon all stages of mealybugs. Mealybug destroyer larvae resemble mealybugs but they are covered with waxy appendages and are more mobile than mealybugs. The mealybug destroyer is more effective from spring through fall than in winter. This predatory beetle prefers temperatures above 60°F (15°C). Release in the evening. Consult with your supplier on recommended release rates. Pest control materials such as dinotefuran (Safari®) and acetamiprid (TriStar®) have been shown to directly harm the mealybug destroyer whereas pyriproxyfen (Distance®) and flonicamid (Aria®) are not directly harmful. Pest Control Materials It is difficult to control or regulate mealybug populations with pest control materials (insecticides) due to their waxy coating, which reduces

contact and penetration of sprays. The young nymphs or crawlers, however, don’t have a waxy covering so they are susceptible to spray applications of many insecticides. Systemic insecticides must be applied preventively while plants are actively growing so that lethal concentrations of the active ingredient are present at feeding sites. Since systemic insecticides do not kill eggs, additional applications may be warranted after 3 to 4 weeks. Contact insecticides such as insect growth regulators, insecticidal soap and horticultural oil, effectively kill young nymphs. However, as eggs hatch throughout the growing season, repeat applications are required. Application frequency depends on the residual activity of insecticides used, varying from 1 to 3 weeks. Thorough coverage of all plant parts is essential. The use of a spreader-sticker may improve coverage and penetration, but may also increase the risk of phytotoxicity (plant injury). Insecticidal soaps and horticultural oils effectively kill eggs, nymphs, and young adults. Systemic insecticides applied as a drench to the growing medium may effectively control or regulate root mealybug populations.

MITES Mites are more closely related to spiders and ticks than to insects. Mites, in general, use their piercingsucking mouthparts (stylets) to puncture plant cells and then withdraw cell contents. All mites are wingless, and their head and thorax are fused together. The body is compact, and oval to oblong in shape. Mites are small (85ºF (29ºC).

Fig. 4.5: Spider mite. A - egg, B - larva, C - protonymph, D - deutonymph, E - adult

Adult females are oval, approximately 1/50 inch (0.5 mm) long and green to orange in color with two dark spots on both sides of the abdomen. Adult females lay up to 12 eggs per day and are capable of laying up to 100 eggs during their 3 -4 week lifespan. Eggs are globular and amber red when viewed with a 10X hand lens. Eggs hatch in approximately three days, and young mite larvae immediately begin feeding. After transitioning through two nymphal stages (deutonymph and protonymph), mites become adults. Females, which do not have to mate to reproduce, begin laying eggs within 1 to 3 days. Scouting Inspect plants weekly for signs of TSM feeding injury. Susceptible plants include angel’s trumpet, bee balm, butterfly bush, cordyline, dahlia, delphinium, Transvaal daisy (gerbera), ivy geranium, mandevilla, New Guinea impatiens, phlox, marigold, salvia, scabiosa, scaevola, thunbergia and verbena. Look for mites on the underside of mature leaves, especially along the midvein and on lower leaves. When scouting, check for the presence of live TSM and their round eggs. Initial infestations commonly occur in warm and dry locations such as near steam pipes, furnaces or heaters, or hanging baskets positioned overhead. Cultural Control Remove weeds in and around greenhouses, as they provide refuge for TSM. Dispose of old plant material including “pet plants” which may harbor populations of TSM. Proper crop irrigation decreases susceptibility to TSM. Avoid overfertilizing plants, especially with soluble forms of nitrogen-based fertilizers, because this promotes the production of succulent growth that is easier for TSM to feed on. Overfertilized plants contain higher concentrations

of amino acids that are essential food sources for TSM, and may enhance development and reproduction of TSM females. Although drip irrigation is a more efficient way to water plants, occasional overhead irrigation will wash TSM off plants. Routine plant inspection helps avoid dealing with TSM populations with all life stages present simultaneously. This in turn maximizes the effectiveness (based on percent mortality) of any miticide applications. Furthermore, this may reduce the number of applications needed, which is important because TSM populations can easily develop resistance to miticides, making control or regulation very difficult. It is essential to rotate miticides with different modes of action in order to delay the onset of resistance. Biological Control Commercially available predatory mites that may control or regulate TSM populations include Phytoseiulus persimilis, Galendromus occidentalis, Neoseiulus (Amblyseius) californicus, Amblyseius (Neoseiulus) fallacis, Amblyseius andersonii, and Mesoseilus longipes. Each species requires and is adapted to different environmental conditions (e.g., temperature and relative humidity). For example, G. occidentalis, M. longipes, N. fallacis, and N. californicus tolerate warmer conditions (>85ºF or 29ºC) and lower relative humidity (30 - 40%) than P. persimilis. Furthermore, these predatory mites generally persist at low populations of TSM. The effectiveness of predatory mites depends on TSM population levels and alternate food sources. For example, P. persimilis only feeds on TSM, while the other predatory mite species either feed on alternative hosts or prey, or on flower pollen. Additional commercially available predators of TSM include the predatory midge, Feltiella acarisuga, and the ladybird beetle, Stethorus punctillum. Consult your supplier for information on release rates. Below are descriptions of the predators commercially available for control or regulation of TSM populations: Mite Predators A.

Phytoseiulus persimilis: This is the most effective predatory mite for control or regulation of TSM populations. Phytoseiulus persimilis spreads among crops to locate TSM

colonies using odors emitted from infested plants. This is a specialist predatory mite that only uses TSM as a food source, feeding on all life stages (eggs, larvae, nymphs and adults). Phytoseiulus persimilis adults are bright red in color, pearshaped with long legs, and are larger and more active than TSM. Adult females lay eggs that are approximately 2 to 3 times as large as TSM female eggs. Both the adults and nymphs actively search plants for TSM. Phytoseiulus persimilis is suitable for use in short-term crops such as bedding plants. Initiate releases early when TSM populations are low or when TSM are first detected. Two to three applications, one week apart, may be required. Make releases near TSM infestations and concentrate releases near localized hot spots. The temperature must be around 68ºF (20ºC) with 75% relative humidity in order for this predatory mite to be effective. Plants need to be touching for the predatory mites to disperse through the crop. Gently sprinkle predatory mites on the lower leaves. When temperatures exceed 86ºF (30ºC), P. persimilis cannot keep up with the reproductive capacity of TSM. When relative humidity is 70°F or 21°C (range: 50 to 85°F, or 10 to29°C) and a relative humidity of 60 to 85%. Neoseiulus (Amblyseius) cucumeris is not compatible with horticultural oil, but populations of the predatory mite may reestablish on plants after spray residues have dried. Spinosad (Conserve®), abamectin (Avid® and other generic products), and mixtures of these two pest control materials may reduce N. cucumeris populations. Mixtures of the fungicide thiophanatemethyl (Cleary’s 3336™ and other generic products) with the pest control materials spinosad (Conserve®) and abamectin (Avid® and other generic products) have been shown to be detrimental to N. cucumeris. C.

Amblyseius swirskii: This above-ground predatory mite feeds on both thripslarvae and whiteflies, and has been shown to effectively control or regulatethrips populations on greenhouse-grown peppers, cucumbers and severalornamental crops. Amblyseius swirskii may also feed on eriophyid mites,broad mite, TSM, and pollen in the absence of prey. Amblyseius swirkskii ismost effective at warmer temperatures (70°F or 21°C) and 70% relativehumidity. It is available in breeding sachets, or can be bulk-applied to thefoliage.

D.

Orius species: These predatory bugs, commonly called ‘minute pirate bugs’, eat immature and adult thrips on leaves and in flowers. As a generalist, it also feeds on aphids and spider mites. Adults are 1/12 to 1/7 inch (2.2 to 3.8 mm) long with flattened bodies, and are typically black and white. Both adults and nymphs are predaceous. Adults tend to reside in open flowers, where adult thrips are usually located. Nymphs may be seen on leaves. Orius is most effective at temperatures of 68 to 85°F (20 to 30° C). Release in the early morning or

late evening; avoid releases in bright sunlight. Minute pirate bugs are most effective on long-term greenhouse vegetable crops, especially those like sweet pepper that produce pollen, compared to short-term crops such as bedding plants. Minute pirate bugs may take up to 12 weeks to establish in a greenhouse so they may be applied to thrips banker plants or habitat plants to encourage early development (see previous section on thrips banker plants). They may also undergo diapause (resting period) during short day conditions. But, several species do not undergo diapause, which means they may be effective during winter conditions. Entomopathogenic Nematodes Steinernema feltiae: Drench applications of this beneficial nematode against fungus gnat larvae may also be effective against thrips pupae in growing media. Successful programs include a drench application to the growing medium followed by weekly spray or sprench applications (see previous section on fungus gnats for application tips). Apply nematodes in the early morning or late evening to avoid desiccation from ultra-violet light, and when thrips mobility is generally slow. Use blackcloth curtains to minimize ultra-violet (UV) light and heat exposure and turn off artificial lights for at least two hours after applying the nematodes. Spray adjuvants may improve application uniformity especially on plants with waxy leaves and allow the nematodes to reach thrips more effectively. Entomopathogenic Fungi A.

Beauveria bassiana: Beauveria bassiana is a naturally occurring entomopathogenic fungus that is commercially available under the following trade names: BotaniGard® and Mycotrol®. In addition to thrips, these materials may provide control or regulation of whiteflies, aphids, and mealybugs. The effectiveness of B. bassiana varies depending on relative humidity levels at the plant surface, life stage (egg, nymph, pupa and adult) of the target insect pest, application rate, crop type, spray coverage, light intensity, season and temperature. For example, insects such as thrips and aphids shed (molt)

their cuticle or exoskeleton. During warm conditions, these insects molt so rapidly that the spores of B. bassiana are shed along with the cuticle so that the entomopathogenic fungus cannot penetrate the insect and initiate an infection. However, the addition of an insect growth regulator such as azadirachtin (Azatin ®, Ornazin®, Molt-X™, AzaGuard, Azahar, AzaDirect, AzaSol) to the spray solution containing B. bassiana may inhibit molting, thus allowing the fungal spores to penetrate the insect cuticle. In addition, applying B. bassiana in early morning may enhance its efficacy. Effective control or regulation of populations of thrips is likely to be achieved when plants are small, when it is easier to obtain thorough spray coverage. Since thrips tend to hide in secluded places, it is essential to thoroughly spray both upper and lower leaf surfaces, and flower and terminal buds. Repeat applications are often needed to maintain low thrips population levels. B.

Isaria (=Paecilomyces) fumosoroseus: This naturally occurring insect fungusis commercially available under the trade names NoFly WP and Preferal.This fungus works best at temperatures of 72 to 86°F (22 to 30°C) andrequires a relative humidity >80%.

C.

Metahizium anisopliae (strain 52): This fungus infects insects on contact andis commercially available as a rice-based granular. It may be used againstthrips pupae in growing media and also as foliar applications against avariety of insect pests.

Pest Control Materials Initiate insecticide applications when thrips populations are low. Thoroughly cover all plant parts including flowers with the spray solution. Insecticides with either contact or translaminar activity are commonly used to control or regulate populations of thrips. Systemic insecticides generally do not move into flowers where adult thrips typically feed; however, they may suppress or even kill nymphs and adults feeding on leaves. Depending on the thrips population, insecticide applications may be required every 3 to 5 days, especially from spring through late fall. Apply insecticides with equipment that produces small spray droplets that are less than 100 microns in diameter

in order to penetrate terminal and flower buds, and open flowers where thrips are usually located. However, in most instances, the use of highvolume application equipment is needed in order to deliver sufficient spray solution to the areas where thrips are located. It is much easier to control or regulate thrips populations when crops are small since this facilitates thorough coverage of all plant parts. Applications to crops with open flowers are generally too late because damage has already occurred. To delay the onset of insecticide resistance it is imperative to rotate insecticides with different modes of action every 2 to 3 weeks (depending on the season) or after more than one generation. Avoid applying insecticides with repellent properties such as the pyrethroid-based insecticides, since this may cause thrips to disperse throughout the greenhouse.

WEEVILS Weevils are the largest group of beetles in the order Coleoptera. They have hardened bodies and distinctly long snouts (mouthparts), and they typically have elbowed antennae. The larvae of many weevil species feed on plant roots while the adults feed on leaves. Many species cannot fly because their wing pads are fused together. The larvae of most species, especially those considered pests of greenhouse-grown crops, are white, grub-like, usually legless and cylindrical. They have well-defined heads and forceful mouthparts. Black Vine Weevils Identification, Biology and Life Cycle

Fig. 4.12: Black vine weevil

A number of weevil species may be encountered in greenhouses, but the black vine weevil (Otiorhynchus sulcatus) is the most important insect pest of greenhouses and nurseries throughout the Northeast. Black vine weevil is 3/8 inch (9.0 mm) long, brown-toblack in color with yellow-brown markings on the wing covers, with ridges extending down the length of the abdomen. The surface of the abdomen has distinct rows of minute punctures. Although black vine weevil adults cannot fly, they can crawl through greenhouse openings. Outdoors, adult female weevils lay eggs in soil or other growing medium near host plants from July through August. Eggs hatch in about two weeks. Larvae are white and legless with yellow-brown heads. When mature, larvae are ½ inch (13.0 mm) long. Black vine weevil larvae feed on the roots of host plants and then overwinter as full-grown larvae. Pupation occurs in cells that they excavate in the growing medium. Emerging adults are all female. Weevil outbreaks may occur when infested nursery stock is introduced into greenhouses. Cultural Control Avoid black vine weevil infestations by buying nursery stock certified weevil-free. Closely inspect incoming plants to exclude weevils from production areas. (Forewings are fused, so adults cannot fly.) Handpick adults if only a few plants are infested in retail setting.

Scouting Although adults are active at night, they may be monitored by placing burlap bands around the base of plant material or positioning pit fall traps in nursery blocks to capture adults walking between plants. Check plant debris and soil or growing medium around plants for the presence of adult black vine weevils. Black vine weevil adults feed at night, notching the edges of plant leaves. During the day, adults stay at the base of host plants such as astilbe, bergenia, epimedium, helleborus, heuchera, heucherella, hosta, physostegia, phlox, primula, saxifraga, sedum and tricyrtis. Larvae feed on roots, causing leaf yellowing, plant stunting and wilting. If plants exhibit these symptoms, check the root system and surrounding root ball for the presence of larvae. Immediately discard heavily infested plants. American willowherb (Epilobium ciliatum), an annual weed in container production, can be used as an indicator plant for black vine weevil adult feeding activity. Biological Control Black vine weevil larvae are susceptible to several biological control agents including fungi and entomopathogenic nematodes. Apply both as drenches to the growing medium to kill larvae. The fungus, Metarhizium anisopliae (Met 52 G) has been demonstrated to control or regulate black vine weevil larval populations in potted plants grown in greenhouses when applied directly to the root system. The most effective entomopathogenic nematode against black vine weevil larvae is Heterorhabditis heliothidis. Both the fungus and the nematode kill young life stages more effectively than older life stages. Commercially available entomopathogenic nematode species for control or regulation of black vine weevil populations include Heterorhabditis bacteriophora and Steinernema kraussei. Pest Control Materials It may be necessary to apply an insecticide drench to protect adjacent plants. Drench applications of the insecticide, bifenthrin (Talstar®) have been shown to effectively kill black vine weevil larvae in containers and in the growing medium. Applications must be made when populations are low. Apply insecticides to potential sources of an infestation in May when adults begin to emerge from the growing medium. The goal is to kill adult females before they lay eggs.

Whiteflies Identification, Biology and Life Cycle The primary whitefly species in greenhouses include the greenhouse whitefly (Trialeurodes vaporariorum) and sweet potato whitefly B-biotype (Bemisia tabaci), which was formally called the silverleaf whitefly (Bemisia argentifolii). Another whitefly species, bandedwinged whitefly (Trialeurodes abutilonia), may enter greenhouses in the fall. None of the whitefly species mentioned here are able to survive outdoors during Northeast winters. Plant material located outside may become infested when winged adults migrate out of greenhouses through openings (e.g., ridge vents, sidewalls and louver vents) as outdoor temperatures increase. Planting infested greenhouse crops outdoors also contributes to infesting outdoor plant material. Whitefly adults, present on weeds or infested host plants outdoors, may migrate into greenhouses in the fall. Nymphs and adults are typically located on the underside of plant leaves. Both nymphs and adults have piercing-sucking mouthparts, which are used to feed on plant fluids. Nymphs may secrete copious amounts of honeydew (clear, sticky liquid) that serves as a growing medium for black sooty mold fungi. Severe infestations of whiteflies may actually defoliate plants. The life cycles of the two whitefly species are similar, consisting of eggs, nymphs, pupae (fourth instar nymph) and adults. Development from egg to adult takes 14 to 40 days, depending on temperature, host plant and whitefly species. Below are descriptions of the common whitefly species found in New England greenhouses. The different species are best distinguished by examining the pupal stage found on the leaf undersides.

Fig. 4.13: Greenhouse whitefly. A - adult, B - egg, C-E nymph, F - Pupa

A.

Greenhouse Whitefly: Greenhousewhitefly adults are more active attemperatures around 75ºF (24ºC).Adults are winged, white, and 1/16inch (2.0 mm) long. Greenhousewhitefly adults hold their wings flat,parallel to the top of the body.Females lay more than 20 eggs in asmall circle. Newly laid eggs arewhite and eventually turn gray.Young nymphs (crawlers) are white,have legs and antennae, and moveshort distances before locatingsuitable places to initiate feeding.More mature nymphs (third and fourth instars) are typically found on thelower leaves. Pupae do not feed, and have distinct visible red eyes.Greenhouse whitefly pupae may possess long waxy filaments encircling theouter edge, and are elevated in profile with vertical sides, resembling“cakes” on leaf surfaces. Sweet potato whitefly B-biotype adults prefertemperatures >80ºF (26ºC).

B.

Sweet potato whitefly. Sweet potato whitefly Bbiotype adults are yellowand smaller than greenhouse whitefly. Their wings are tilted, and held rooflike over their bodies. Adult females live up to six weeks, and produce upto 200 eggs, which are randomly laid in small clusters on new plant growth.Newly laid eggs are white and then turn amber-brown. Young nymphs(crawlers) have legs and antennae, and move short distances before locatingsuitable places to initiate feeding. More mature nymphs (third and fourthinstars) are typically found on the lower leaves. Sweet potato whitefly B-biotype nymphs are yellow, oval and dome-shaped, and do not possess long waxy filaments. In addition to the two whitefly species described above, a new biotype of B. tabaci, the Q-biotype, was reported in the U.S. in 2006. This biotype may be problematic because it is known to be resistant to a number of commonly used insecticides including imidacloprid (Marathon®), thiamethoxam (Flagship®), acetamiprid (TriStar™), and the insect growth regulators

buprofezin (Talus®) and pyriproxyfen (Distance®). This biotype also vectors leaf curl virus of tomatoes. Submit specimens or samples to specialists for genetic testing because it is difficult to determine which biotype is present based on visual inspection. Winged adults emerge in 1 to 2 weeks through Tshaped apertures on the top of pupal cases, which remain attached to leaf surfaces. Empty pupal cases may be present on old or senescing leaves, and may be mistaken for live whitefly nymphs. Adult females initiate egg-laying 2 to 3 days after emerging. C.

Bandedwinged whitefly: Bandedwinged whitefly is not an important whitefly species in New England greenhouses. However, adults may be captured on yellow sticky cards in the fall and cause concern among greenhouse growers. In the North, this whitefly species can only survive the winter inside greenhouses. Outdoors, bandedwinged whitefly may be found on weeds such as ragweed (Ambrosia species) and velvetleaf (Abutilon theophrasti). Adults are yellow in color with a green tinge on the thorax. The front wings are marked with two zigzag, smoky gray bands. When the wings are folded over the body, these lines appear to be continuous from wing to wing. Hind wings lack bands. Waxy filaments encircle the outer edge of fourth instar nymph (pupae) bodies, and a distinct darkened line extends down the center of the body. This trait helps distinguish bandedwinged whitefly from other whitefly species.

Fig. 4.14. Silverleaf whitefly. A - Adult, B - Egg, C - Crawler, D - Nymph

Scouting Visually inspect incoming plant material for the presence of whiteflies. Once a crop is established, monitor weekly by checking the undersides of 1 to 2 leaves on 10 to 20 plants throughout the greenhouse to detect eggs, nymphs and pupae. Use yellow sticky cards to capture whitefly adults. Put sticky cards just above the crop canopy and replace weekly, or use one side one week and the other side the next week so as to use a single sticky card for two weeks (overlay the unused side with the sticky cover). Raise sticky cards as plants grow. As a general rule, use one sticky card per 500 to 1,000 ft2 of greenhouse space. Use more sticky cards for susceptible crops. Count the number of adult whiteflies captured on sticky cards weekly and record this information in a computer database. Use this information to decide if natural enemy releases or insecticide applications are needed. Cultural Control Avoid overfertilizing crops as this increases their attractiveness to adult whiteflies. Whiteflies may be introduced into greenhouses on infested cuttings or plants arriving from outside sources. Residual or carryover plants from previous crops and stock plants may also be sources of whiteflies. Using appropriate sanitation practices like weed removal helps alleviate whitefly problems in subsequent cropping cycles. Biological Control

Two parasitoid species commercially available for controlling or regulating whitefly populations in greenhouses are Encarsia formosa and Eretmocerus eremicus. Below are descriptions of these parasitoids. Whitefly parasitoids A.

Encarsia formosa: This parasitoid, commercially available since the 1970s, iseffective against the greenhouse whitefly, particularly in long-term (morethan four months) crops such as tomatoes and cucumbers. Encarsia formosais less effective on tomatoes with hairs or trichomes because the hairs inhibitthe parasitoids’ ability to detect whiteflies, and adult females to lay eggsin whitefly nymphs. Adult female parasitoids may also feed on youngnymphs. Females lay eggs in nymphs; larvae emerge from eggs andconsume the internal contents of the whiteflies. Larvae eventually pupate,and emerging adults create a circular hole with their mouthparts, which isused to exit the parasitized whitefly. Parasitized whiteflies are typicallyblack. Encarsia formosa is most effective at 70 to 80ºF (21 to 26ºC), and 50 to80% relative humidity. Adults don’t fly when ambient air temperatures arebelow 65ºF (18ºC) and survival is reduced at temperatures >86ºF (30ºC). Remove yellow sticky cards before releasing E. formosa in order to avoid capturing adults on the cards. Replace sticky cards 3 to 4 days following release. Most E. formosa are commercially available as pupae glued to paper cards. Suspend the cards in the lower canopy of plants to avoid desiccation from direct sunlight. Adults emerge from the pupae and fly upward. Introduce cards weekly starting when whiteflies are first detected. In general, for most crops, continue making releases until 80% of the whitefly population has been parasitized. When scouting, look for the distinct black parasitized greenhouse whitefly pupae. Encarsia formosa is very sensitive to direct sprays and even dried residues of products including pyriproxyfen (Distance®), spinosad (Conserve®), chlorfenapyr (Pylon®), acetamiprid

(TriStar™) and pyridaben (Sanmite®). B.

Eretmocerus eremicus: This parasitoid has been commercially available sincethe 1990s for control or regulation of the Bbiotype of Bemisia tabaci onpoinsettia. It tolerates warm temperatures since it is native to desert areasof California and Arizona. Besides directly parasitizing whitefly nymphs, E. eremicus adult females kill nymphs by host feeding, which may actually maintain whitefly populations at low levels. Eretmocerus eremicus attacks both sweet potato whitefly Bbiotype and greenhouse whitefly. Eretmocerus eremicus is shipped as pupae that are either glued to paper cards or in blister packs or in bottles. Do not release in direct sunlight. Contact biological control suppliers for information on release rates. Prior to release, remove yellow sticky cards, which attract and capture the emerging parasitoids. Replace yellow sticky cards four days after releases have been made. Development and activity are optimized at 77 to 84ºF (25 to 29ºC). E. eremicus is inactive at temperatures >86ºF (30ºC). When scouting, look for parasitized whiteflies. Greenhouse whitefly pupae are yellow while sweet potato whitefly Bbiotype pupae are yellow-brown. In general, E. eremicus is more tolerant of exposure to pest control materials than E. formosa. For long-term crops (5 months or longer) infested with greenhouse whitefly, E. formosa releases are more efficient because it reproduces better on greenhouse whitefly than E. eremicus, which tends to act as a predator by hostfeeding on the nymphs. Once E. formosa populations establish in greenhouses, only occasional releases may be needed. Use of E. formosa for control or regulation of greenhouse whitefly populations is recommended for long-term plantings such as conservatories; greenhouse-grown vegetables such as tomatoes, peppers and cucumbers, whose production time exceeds 20 weeks; or crops such as cut flowers since the foliage is not sold. In short-term floral crops (less than four months) with lower

whitefly thresholds, or crops like poinsettia, E. eremicus is recommended. Releases must begin when plants are potted or when they emerge as seedlings, and should continue during the cropping cycle. Whitefly Predators A.

Amblyseius swirskii feeds on whitefly eggs and nymphs as well as thrips. Thispredatory mite is most effective at warmer temperatures (70ºF or 21ºC) anda relative humidity of 70%. It is available in breeding sachets, or in bulk tobe applied to plant leaves. Consult with supplier for release rateinformation.

B.

Delphastus pusillus is a small (1.3 to 1.4 mm) long dark brown-to-blackpredatory beetle that attacks all stages of whiteflies, but prefers eggs andnymphs. Delphastus adults and larvae are predacious. Optimumtemperatures are 75 to 80°F (24 to 27°C), and adults do not fly attemperatures below 55°F (13°C). Release in the early morning or evening.Consult with your supplier on release rates. Delphastus avoids feeding onparastisized whiteflies, so is compatible with the parasitic wasps describedabove.

Entomopathogenic Fungi A.

Beauveria bassiana: This fungus is commercially available for control orregulation of whitefly populations. Applications must be initiated beforewhitefly populations are high. The fungus works best if the relativehumidity is >90%.

B.

Isaria (=Paecilomyces) fumosoroseus: This fungus is available under the tradenames NoFly WP and Preferal. It attaches to whitefly eggs, nymphs, pupaeand adults. It requires a relative humidity of 68 to 100% and temperaturesof 72 to 86°F (22 to 30°C). Foliar applications of Isaria have been successfullycombined with releases of Encarsia formosa in commercial greenhousetomato production.

Pest Control Materials

The two application methods for pest control materals (insecticides) to control or regulate populations of whiteflies on greenhouse-grown crops are (1) foliar applications directed at adults and nymphs; and (2) systemic insecticides applied to the growing medium to control or regulate nymphal populations. Whitefly eggs and pupae tolerate most insecticides whether used as sprays or drenches, but horticultural oils kill all the life stages if thorough coverage is achieved. Similarly, when using contact insecticides, it is imperative to thoroughly cover leaf undersides. Repeat applications may be needed over a 2 to 3 week period, depending on the extent of the whitefly population and residual activity of the insecticide. Translaminar insecticides may control or regulate whitefly populations for extended periods of time. Start applying systemic insecticides, such as neonicotinoids, when plants are actively growing to increase uptake of the active ingredient. After systemic insecticide application, it may take several days to weeks before whitefly populations start to decline. This may vary, depending on the water solubility of the systemic insecticide, plant age and growing medium. The use of insect growth regulators may be preferable early in the cropping cycle since many of these insecticides are compatible with whitefly parasitoids.

ADDITIONAL PESTS Millipedes Millipedes are not insects, but belong to the class Diplopoda. They may feed on decaying plant material and may sometimes feed upon young seedlings’ roots, especially in ground or soil bed production. Most species curl into a coil when disturbed. They have two pairs of legs per body segment, and short antennae. Springtails Springtails are very small (1/5 inch or 5 mm long), wingless insects, and are brown to purple. They have a distinct spring-like apparatus (furcula) at the tip of their abdomen that allows them to jump. They are commonly seen on or in moist growing medium in containers, especially if plants are overwatered. Springtails may be found on potato disks used to monitor for fungus gnat larvae. They are sometimes misidentified as thrips. Springtail

females lay eggs in growing medium or soil underneath benches. As they mature, “nymphs” change color and grow. Springtails are primarily scavengers, feeding on decaying organic matter, algae or fungi. However, some species prey on other springtails or nematodes. In general, springtails are not considered a greenhouse pest because they do not feed on plants. Slugs and Snails Identification, Biology and Life Cycle Slugs and snails are not arthropods but are classified as mollusks, closely related to oysters and clams. Snails have hard-shell coverings that protect their bodies. Slugs do not have an outer hardened covering, but they are covered with a copious amount of mucous-like slime that protects their bodies from desiccation. Slugs and snails vary in length from ¾ to 1½ inches (2 to 4 cm). Their color ranges from pale yellow to lavender to purple. Slugs and snails lay translucent pearl-shaped eggs in clusters of 20 to 100 in cool, moist locations such as in soil or growing medium, underneath mulch, boards, and/or plant pots. Eggs hatch in less than 10 days at 50ºF (10ºC). Young slugs and snails resemble adults but are lighter in color and smaller. They mature in 3 to 12 months, and adults may live a year or more. Slugs contain both male and female organs, and may alternate sexes during adulthood. Self-fertilization is also possible. Scouting Inspect damp, moist areas for slugs and their egg clusters that are covered with a gelatinous shell giving them a somewhat milky appearance. Slugs and snails feed on a wide range of greenhouse-grown crops at night. They use their chewing mouthparts to create holes in leaves and stems. Feeding damage from slugs and snails may be misdiagnosed as that of caterpillars. However, caterpillars typically leave fecal deposits on plant leaves and stems. Also, slugs and snails completely eat leaves and stems while caterpillars may leave portions of stems, leaf veins or the epidermal layer untouched. Slugs and snails also leave shiny mucous-like slime trails in areas they have visited. These trails may be more evident when slugs are most active in the evening or early morning hours or after plants are irrigated. Cultural Control Slugs and snails are commonly introduced into greenhouses via infested

plant material, unsterilized growing medium (soil and sand) or supplies (flats and (containers) that have been stored outdoors for an extended time period, especially if perched on soil. Sanitation practices such as weed removal in and around the greenhouse and removing debris (boards and old flats) helps reduce problems with slugs and snails. Sterilizing growing medium and thoroughly cleaning containers or flats prior to use may also alleviate slug and snail problems. Slugs and snails tend to avoid crossing copper barriers, so encircling bench posts with strips of copper sheets may prevent both pests from reaching plants on benches. Pest Control Materials Pest control materials (molluscicides) or baits that are used to control or regulate populations of slugs and snails may contain three active ingredients: metaldehyde (Deadline®), methiocarb (Mesurol®), and iron phosphate (Sluggo®). Metaldehyde does not directly kill slugs and snails, but causes paralysis, resulting in secretion of copious amounts of mucous. Slugs and snails become immobile and desiccate. Metaldehyde is most effective on warm, sunny days. In cool, moist conditions, slugs and snails may actually recover from exposure to metaldehyde. Metaldehyde is very sensitive to environmental conditions and decomposes rapidly when exposed to direct sunlight, but newer formulations appear to resist breakdown when exposed to sunlight. The nerve toxin methiocarb interferes with nerve-impulse transmission. It also acts as a stomach poison, which means slugs and snails must consume the material in order to be affected. This material is effective in cool, moist conditions. Both metaldehyde and methiocarb are toxic to dogs and cats. Iron phosphate, a stomach poison, contains an attractant and the heavy metal, iron. Iron is toxic to slugs and snails, reducing their mobility and causing eventual death. Iron phosphate is not toxic to dogs and cats. Sowbugs Sowbugs are not insects; they belong to the order Isopoda. Sowbugs are commonly found under containers or flats in moist areas such as propagation greenhouses, because they cannot control the loss of moisture from their bodies. They eat mostly decaying organic matter, although they do feed on young plant roots. Sowbugs have a distinctive “armored” appearance, and often roll up into a ball when disturbed. To reduce sowbug numbers, remove

weeds, growing medium and plant debris, and other debris like boards from inside and around the greenhouse. Symphylans

Fig. 4.15: Symphylan

Symphylans are not insects since they belong to the class Symphyla. They are 1/25 to 1/3 inch (1 to 8 mm) long, slender, wingless, white arthropods with 15 to 20 body segments and long antennae. Adults are 3/8 inch (9 mm) long. They resemble centipedes, but they are not predaceous like centipedes. Instead they feed on plant roots and seeds in the growing medium. Extensive or high populations can stunt growth and even kill plants. To determine the presence of symphylans, remove a plant (along with its roots) from the container, place the root ball into a receptacle of water and swirl the contents for several minutes. Any symphylans present will rise and float on the water surface. There is minimal information bout control or regulation of symphylan populations. However, avoid overwatering plants in order to create an environment that is not conducive for development of symphylan populations.

5: Fungal and Bacterial Diseases INTRODUCTION Most diseases affecting greenhouse-cultivated crops are caused by one or more fungal or bacterial infestations. Some fungi and bacteria primarily damage the root system (such as the fungi Fusarium, Pythium, and Phytophthora and the bacterium Agrobacterium rhizogenes), while others primarily manifest above ground (like the fungus Phytophthora infestans or bacteria like Clavibacter michiganensis subsp. michiganensis). Bacteria cannot be combated using crop protection agents, so solutions have to be found in prevention and in competition with other bacteria. Unlike bacteria, fungi can be treated using crop protection agents. However, public opinion and the market are increasingly insisting on a residue-free product, making the use of crop protection agents against fungi more problematic. Fungal leaf spots are common on many vegetable plants. Experience and regular monitoring will alert gardeners as to the seriousness of the problem. Foliar diseases are frequently weather dependent and vary in severity from season to season according to rainfall and temperature. In some cases leaf spots will not spread or cause much damage, but in other cases management actions should be taken promptly because the disease will continue to spread, perhaps defoliating the plant and reducing yields and eating quality. The first step is to learn about the common vegetable crop diseases in your area and, when disease symptoms are observed, identify the disease and determine the least-toxic management strategy. Early blight (Alternaria solani) of tomato is an example of a foliar disease that appears with great frequency in the northeast and mid-Atlantic states and has the potential to defoliate plants during the growing season. Powdery mildew, on the other hand, is a fungal disease that usually infects pumpkins and squash late in the growing season and is not typically a serious

problem for backyard gardeners. Bacteria are single celled, microscopic organisms, bounded by a cell wall, that cause plant diseases. Bacteria are much smaller than fungi but can cause severe symptoms. Bacterial pathogens can cause soft rots of fruits, vascular wilts (e.g. bacterial wilt of cucumber and muskmelon), and leaf spots and blights (e.g. bacterial spot of pepper, bacterial blight of peas).

FUNGAL DISEASES Fungi are organisms that are classified in the Kingdom “Fungi”. They lack chlorophyll and conductive tissue. Until a few years ago, fungi were considered lower forms of plants, but today are classified as a group by themselves. Because fungi cannot manufacture their own food (due to lack of chlorophyll), they must obtain it from another source as either a saprophyte or parasite. Most fungi encountered are saprophytic (feed on decaying organic matter). The parasitic fungi, those that derive their sustenance from living plants, are the group of interest in plant health. In dry climates like Colorado, fungi are the most frequently causes of plant diseases. A fungus “body” is a branched filamentous structure known as mycelium. One single thread is called a hypha (hyphae, plural). Most fungi reproduce by spores, reproductive structures that unlike seeds contain little stored food. Spores are the main dispersal mechanism of fungi and can remain dormant until germination conditions are appropriate. Many fungi over-winter as fruiting structures embedded in dead plant tissue. When a spore comes in contact with a susceptible plant, it will germinate and enter the host if the proper environmental conditions are present. Hyphae develop from the germinated spore and begin to extract nutrients from host plant cells. The hyphae secrete enzymes to aid in the breakdown of organic materials that are ultimately absorbed through their cell walls. Fungi damage plants by killing cells and/or causing plant stress. Fungi are spread by wind, water, soil, animals, equipment, and in plant material. They enter plants through natural openings such as stomata and lenticels and through wounds from pruning, hail, and other mechanical damage. Fungi can also produce enzymes that break down the cuticle, the outer protective covering of plants.

Fungi cause a variety of symptoms including leaf spots, leaf curling, galls, rots, wilts, cankers, and stem and root rots. Fungi are responsible for “damping off” symptoms associated with seedlings.

DAMPING OFF Damping off is the fungal infection of seeds or seedlings that leads to death. When infected with damping off, seeds may fail to germinate. In other situations, seedlings develop but eventually fall over and die. An examination of stems at the soil line reveals a discolored, “pinched in” appearance. Most plants are susceptible to damping off because of the soft immature nature of seedling tissue that is more susceptible to infection. The best method to manage damping off is to avoid it in the first place. For starting seeds indoors, use pasteurized soil or planting mix and ensure that plants receive optimum light, water, and heat for rapid germination and growth. In home situations, damping off frequently develops due to poor lighting and overwatering. These conditions stress plants and make conditions optimal for the development of the soil-borne organisms that cause damping off. In the garden, plant seeds at appropriate times (soil temperature for rapid germination) for the crop and avoid overwatering for optimal germination and growth. A strong healthy plant is better equipped to fight off infection. Scientists continue to study the role of hyperparasites (parasites of parasites) in disease management. Several biological pesticides have been developed from naturally occurring hyperparasitic fungi and bacteria. The organisms protect plant roots against invasion by harmful soil pathogens. These biological pesticides must be applied prior to the development of damping off so the beneficial organisms have time to grow and colonize roots. They cannot be used as “rescue” treatments.

LEAF SPOTS One of the most common fungal plant symptoms is leaf spotting. Leaf spot symptoms are caused by many different fungi. Generally, fungal leaf spots possess a distinct dark brown or red margin between the interior (dead)

and exterior (healthy green) tissue called aborder. [Fig. 5.1] Fungal fruiting structures (reproductive structures) are usually embedded in the dead interior. Frequently a “halo” of yellow or red color develops around the border. A halo indicates recently killed tissue that will eventually die. Because of the cycle of killing tissue and creating a border, then killing more tissue and creating another border, many fungal leaf spots take on a target-like appearance.

Fig. 5.1. Fungal Leaf Spot

To confuse matters, a series of drought events can cause damage that exhibits alternating light and dark bands. Additionally, fruiting structures may not be obvious in dry climates like Colorado. To positively identify a fungal leaf spot, it is best to either culture tissue from the sample or look for spores under a compound microscope. Example of common leaf spot diseases in Colorado include Marssonia and Septoria Leaf Spots of cottonwoods and aspen, Ink Spot of Aspen, and Early Blight of tomatoes and aspen. For additional information, refer to the following. • • • •

Aspen and Poplar Leaf Spots, Aspen Leaf Spot, Marssonina Leaf Spot Sycamore Anthracnose.

ASPEN AND POPLAR LEAF SPOTS Foliage diseases can reduce the aesthetic value of aspen and cottonwood. Occasionally, a severe disease outbreak causes premature defoliation or dieback of parts of the tree.

If a tree loses its leaves early in the season, it may grow new ones and its health is not seriously affected. If it loses them in midsummer, however, growing new leaves may prevent the tree from fully hardening off before cold weather or reduce the amount of stored food. This leads to increased danger of frost damage, reduced growth, and predisposition to other diseases or insects. If the tree loses its leaves late in the season, it will not grow new ones or lose much vigor. Marssonina Leaf Spot The fungus Marssonina causes the most common foliage disease on aspen and cottonwoods in urban and forested areas of Colorado. Marssonina leaf spots are dark brown flecks, often with yellow halos (Fig. 5.2). Immature spots characteristically have a white center. On severely infected leaves, several spots may fuse to form large black dead patches in late summer (Fig. 5.3). Spots also may develop on leaf petioles and succulent new shoots.

Fig. 5.2: Marssonina leaf spot on cottonwood

Fig. 5.3: Marssonina leaf spot on aspen, late symptoms

Marssonina survives the winter on fallen leaves that were infected the previous year. With spring and warmer, wet weather, the fungus produces microscopic spores that are carried by the wind and infect emerging leaves. Early infections are rarely serious, but if the weather remains favorable, spores from these infections can cause a widespread secondary infection. Heavy secondary infections become visible later in the growing season and cause premature leaf loss on infected trees. Septoria Leaf Spot The fungus Septoria causes a common foliar disease mainly on cottonwoods and occasionally aspen in urban areas .

Fig. 5.4: Septoria leaf spot in late summer showing irregular brown to black spots.

The appearance of Septoria leaf spots varies considerably between tree species and with time. Symptoms include, in early summer, a distinct tan circular spot with black margins and small black fruiting bodies in the center, and finally in late summer, irregular brown to black spots that coalesce into large areas (Fig. 5.4).

The disease is rarely a problem on plains and eastern cottonwoods but can cause defoliation at least a month early and visual quality loss on lanceleaf cottonwoods. In wetter climates, another species of the fungus also causes cankers on twigs and main stems. Septoria survives the winter on fallen leaves that were infected. With spring and warmer, wet weather (70 to 75° F), the fungus produces microscopic spores that are carried by the wind and infect emerging leaves. Early infections are rarely serious. If the weather remains favorable, spores from these infections can cause a widespread secondary infection. Heavy secondary infections become visible later in the growing season and cause premature leaf loss on infected trees. Ink Spot of Aspen The fungus Ciborinia causes a leaf disease of aspen commonly known as ink spot. The first symptoms of ink spot appear in late spring to early summer as tan to brown areas on the upper leaf surfaces. Concentric, discolored ring patterns may become visible as the fungus advances through the leaf (Fig 5.5). These concentric patterns can be confused with leafminer attacks. Infected leaves may be totally brown by midsummer, while adjacent uninfected leaves remain green. Raised black bodies begin to appear on affected brown leaves. These hard masses of fungal material (sclerotia) are oval and nearly ¼ inch long. These are the “ink spots” that give the disease its common name. In late summer, these spots may fall out, leaving a characteristic “shot hole” effect on leaves that remain on the tree. This disease is especially prevalent in dense aspen stands. Early season defoliation may reduce growth.

Fig. 5.5: Ink spot disease on aspen in early summer.

The sclerotia that fall from infected leaves are the overwintering stage of the fungus. Wet spring weather stimulates fruiting bodies to form from the sclerotia and produce spores. Spores are blown and splashed from the ground to developing leaves. Leaf and Shoot Blight Leaf and shoot blight, caused by the fungus Venturia, is a disease affecting young aspen and cottonwood tissue primarily in the mountains.

Fig. 5.6: Shoot blight on aspen showing dark and curled stem

In the spring, symptoms first become visible on leaves near shoots infected the previous season. Brown to blackened, irregularly shaped areas spread through the leaves, causing them to dry and become distorted. Typically, the fungus spreads down through the succulent new shoot, causing cankers that blacken and curl the not yet strengthen (lignified) stem tip until it resembles a shepherd’s crook (Fig. 5.6). Death of new shoots causes distorted, shrubby growth. The leaf and shoot blight fungus survives the winter mainly on shoots infected the previous season. Spores are windblown early in the season and infect newly expanding leaves and shoots. As the season progresses, uninfected tissue becomes more resistant to the disease. Leaf Rusts

Fig. 5.7: Leaf rust.

A rust disease caused by the fungus Melampsora is often seen on aspen

and cottonwood. Though common, this disease rarely causes serious damage to trees since the disease develops late in the summer/fall and rarely causes early defoliation. The disease damages leaves after most photosynthetic needs for the tree are completed. The disease is easily recognized by small, yellow-orange pustules that are scattered on the lower leaf surfaces (Fig 5.7). These orange pustules are most visible in late summer and early fall. The life cycle of this fungus requires two different tree hosts. During wet spring weather, spores are released from the fungus, which has overwintered on fallen cottonwood or aspen leaves. These spores infect evergreen needles, such as Douglas-fir, pine, fir or spruce, where they cause little damage. After two to three weeks, spores are produced on these evergreen hosts and are blown to aspen or cottonwood leaves. Once the rust is established on aspen or cottonwood hosts, it can multiply rapidly under favorable wet conditions throughout the summer. Several years of heavy infections can cause some growth losses, especially on younger trees. Fallen infected leaves shelter the fungus until the next year’s disease cycle. Disease Management Tree resistance is the best way to prevent foliar diseases. Several poplar hybrids or species are resistant to one or more of these diseases. Ask your local nursery for a resistant variety. Some aspens are resistant to leaf spots, but aspen production methods make it difficult to select trees for resistance. Sanitation is an effective control for some foliar diseases. Fall removal of infected leaves, twigs and branches can reduce the amount of disease the next spring. Raking and destroying infected leaves can reduce Marssonina and Septoria leaf spot, ink spot and leaf rust. The shoot blight fungus overwinters in diseased stems and twigs, so it can be pruned out to reduce new infections. Keep leaves as dry as possible to reduce the incidence of leaf spots: • Irrigate in early morning so leaves can dry out. • Keep sprinkler patterns adjusted so leaves don’t stay wet. • Space trees apart to reduce humidity to help prevent leaf diseases. Fungicide sprays are not normally needed but if applied early enough, can prevent foliage diseases. Spraying will prevent only new infections; it will not cure leaves already infected. If an infection is developing on

particularly valuable trees, or if there is good reason to believe an infection is imminent, the trees can be sprayed with fungicides. Trees that perennially have foliar diseases should be sprayed at bud break and then two or three times during the growing season at 12-to 14-day intervals. Check fungicide labels carefully since allowable uses and rates can change. Fungicides labeled for leaf spots should work well for all these diseases except for the rusts since they are in a different fungal group. Other fungicides may be required for rust diseases. Please follow label rates and directions when applying.

ASPEN LEAF SPOT Aspens are susceptible to many problems in the urban landscape. One of those problems is aspen leaf spot, a disease caused by fungus. Aspen leaf spot, or Marssonina leaf spot, is common on aspen trees in the late summer and autumn. A previous spring of warm, rainy conditions creates the perfect environment for the disease to develop and spread. When affected by the disease, black spots form on aspen leaves. However, black spots may develop for a variety of reasons, so it’s important to recognize all of the symptoms of aspen leaf spot. Symptoms occur between July and frost. Leaves affected with the disease will be scattered randomly throughout the tree and spots will appear randomly on the leaves. The spots are dark brown to black circles containing rings, looking almost like a bullseye. Sometimes the edges of the spots may appear irregular or feathery.

Fig. 5.8. Aspen Leaf Spot

MARSSONINA LEAF SPOT

Fig. 5.8. Marssonina leaf spot

Marssonina leaf spot is a fungal disease of aspen, poplar and cottonwood. Infection begins in the spring at bud break when spores are blown or splashed to the tender, newly-developing leaves. Brown spots with yellow halos begin to appear on leaves in mid to late July. Many of these spots eventually merge together, creating large brown blotches. Leaves may drop prematurely. To manage the disease, rake up fallen leaves in the autumn and dispose or compost. If there is a history of severe infestations, the fungicides Daconil may be used in the spring beginning at budbreak. Fungicides are of no value by the time symptoms appear. The disease does not kill trees, but stresses them and may weaken them over time.

SYCAMORE ANTHRACNOSE Anthracnose is the common name given to a group of fungal pathogens which cause dark, usually sunken lesions. The term anthracnose is from the Greek word for coal or charcoal. These are typically diseases of leaves, stems or fruits. The sycamore anthracnose fungal organism attacks sycamore trees early in the spring causing a rapid wilt of newly emerging leaves. This rapid wilting is frequently misidentified as frost damage. Larger, more mature leaves develop a brown growth along the main veins. Infected leaves often curl and eventually fall, littering the ground. The fungus involved is Apiognomonia errabunda & veneta (synonyms = Gnomonia errabunda & veneta); anamorphs = Discula umbrinella & platani. The sycamore leaf is naturally fuzzy. Do not confuse this natural fuzziness with infection by anthracnose fungus.

Cankers Cankers often form on the twigs and branches at the base of blighted leaf clusters. Cankers restrict water and nutrient movement to the leaves resulting in twig die-back, chlorosis and scorch on leaves, and even kill larger branches. These cankers are active during wet cool springs and produce spores that spread to neighboring twigs, leaves, and other sycamore trees.

Fig. 5.10: This sunken dead area (canker) is due to Anthracnose

Fig. 5.11: The ‘witches broom’ is where numerous shoots grow from the same point.

Cankers also develop in larger branches, girdling and eventually killing them. Small, black fruiting bodies of the causal fungus appear in the discolored bark of dead twigs and branches. Repeated annual killing of twigs results in clusters of old dead twigs and live branches creating what are called “witches’ brooms.” Weather Influences Weather determines the severity of anthracnose. Frequent rains and cool temperatures promote the disease. If the average temperature during the twoweek period following emergence of the first leaves is below 55 degrees F,

the shoot-blight phase of the disease will be serious. Disease intensity decreases as the average temperature increases from 55 to 60 degrees. Little or no anthracnose will occur if average temperatures during this susceptible stage are above 60 degrees. Cultural Practices Gather and destroy all fallen leaves and twigs. They will produce fungus spores the following spring if not destroyed. Prune out all infected twigs and branches and destroy them. Cut out cankers in large limbs to reduce reinfection. Remove the dead, cankered tissue down to healthy wood. Dry winters weaken trees, increasing the effects of diseases. To reduce this problem, water trees once a month during snowless winters. Water when air temperatures are above freezing and early enough during the day to allow water to soak in before nightfall. Resistance The American sycamore (Platanus occidentalis L) is much more susceptible to anthracnose than the London (Platanus x acerifolia) and Oriental plane (Platanus orientalis). Due to this high susceptibility, planting of the American sycamore should be avoided. The Oriental plane, a shorter, less graceful tree, is highly resistant to anthracnose but rarely grown in the United States. The London plane tree (P. x acerifolia, synonyms P. x orientalis or P. hybrida) is a cross between the Oriental plane (P. orientalis) and the highly susceptible American sycamore (P. occidentalis). The London plane cultivars, ‘Bloodgood’, ‘Columbia’ and ‘Liberty’ are resistant to anthracnose and are good choices for planting where the sycamore anthracnose fungus is a problem.

MILDEW Powdery Mildew is one of the most common diseases in dry climates like Colorado. General symptoms include a white or grayish powdery growth on leaves. It thrives in warm dry climates, often explodes in small yards with limited air movement, and in the fall as nighttime humidity rises [Fig. 5.12].

There are many species of mildew fungi, each being host specific. In Colorado for example, it is common on ash, lilac, grapes, roses, turfgrass, vine crops (cucumbers, melons, and squash), peas, euonymus, cherry, apple, crabapple, pear, Virginia creeper, and others.

Fig. 5.12. Powdery Mildew on lilac.

Management is centered on a variety of cultural techniques. Avoid crowing plants as the lack of air circulation favors powdery mildew. Select resistant varieties where possible. Avoid late-summer application of nitrogen fertilizer as it may pushes growth of tender young leaves that are more prone to mildew. Avoid overhead irrigation as it raises relative humidity. Removed and destroy infected plant parts. A variety of fungicides found in the home garden trades are effective on powdery mildew. Symptoms Even though there are several types of powdery mildew fungi, they all produce similar symptoms on plant parts. Powdery mildews are characterized by spots or patches of white to grayish, talcum-powder-like growth. Tiny, pinhead-sized, spherical fruiting structures that are first white, later yellowbrown and finally black, may be present singly or in a group. These are the cleistothecia or overwintering bodies of the fungus. The disease is most commonly observed on the upper sides of the leaves. It also affects the bottom sides of leaves, young stems, buds, flowers and young fruit. Infected leaves may become distorted, turn yellow with small patches of green, and fall prematurely. Infected buds may fail to open. Conditions That Favor the Disease The severity of the disease depends on many factors: variety of the host plant, age and condition of the plant, and weather conditions during the

growing season. Powdery mildews are severe in warm, dry climates. This is because the fungus does not need the presence of water on the leaf surface for infection to occur. However, the relative humidity of the air does need to be high for spore germination. Therefore, the disease is common in crowded plantings where air circulation is poor and in damp, shaded areas. Incidence of infection increases as relative humidity rises to 90 percent, but it does not occur when leaf surfaces are wet (e.g., in a rain shower). Young, succulent growth usually is more susceptible than older plant tissues. About the Fungi Powdery mildews are host specific - they cannot survive without the proper host plant. For example, the species Uncinula necator, which causes powdery mildew on grape and linden, does not attack lilac. Similarly, Microsphaea alni affects elm, catalpa, lilac and oak but not turfgrass. Powdery mildews produce mycelium (fungal threads) that grow only on the surface of the plant. They never invade the tissues themselves. The fungi feed by sending haustoria, or root-like structures, into the epidermal (top) cells of the plant. The fungi overwinter on plant debris as cleistothecia or mycelium. In the spring, the cleistothecia produce spores that are moved to susceptible host tissue by splashing raindrops, wind or insects. Control Cultural Several practices will reduce or prevent powdery mildews. Many plants, such as roses, vegetables and Kentucky bluegrass, have cultivars, which have been developed to be resistant or tolerant to powdery mildew. Inquire about resistant varieties before a purchase. If resistant varieties are unavailable, do not plant in low, shady locations. Once the disease becomes a problem: •

Avoid late-summer applications of nitrogen fertilizer to limit the production of succulent tissue, which is more susceptible to infection. • Avoid overhead watering to help reduce the relative humidity. • Remove and destroy all infected plant parts (leaves, etc.). For infected vegetables and other annuals, remove as much of the plant and its debris

in the fall as possible. This decreases the ability of the fungus to survive the winter. Do not compost infected plant debris. Temperatures often are not hot enough to kill the fungus. • Selectively prune overcrowded plant material to help increase aircirculation. This helps reduce relative humidity and infection.

CANKERS Cankers are discolored, sunken areas found on plant stems, branches, and trunks. They damage plants by killing the conductive tissue. Cankers may be caused by fungi, bacteria, virus, and abiotic disorders such as sunscald and hail. [Fig. 5.13]

Fig. 5.13: Powdery Mildew on lilac.

Fungal cankers contain fruiting structures embedded in the discolored canker. Plants with cankers may exhibit branch dieback, leaf loss, and/or poor growth above the damaged area. Common fungal cankers Cytospora (Cytospora sp.) and Thyronectria (Thyronectria sp.). Common bacterial cankers in Colorado include fireblight (Erwinia amylovora). For additional information, refer to the following : • • • •

Canker Diseases on Deciduous Cytospora Canker Fire Blight Honeylocust Diseases.

Canker Diseases on Deciduous Trees Aspen, cottonwood, mountain ash, willow and elm trees with environmental or weather-related stress are often more susceptible to canker

diseases. These diseases also strike thin-barked trees that are susceptible to sun burns.

Fig. 5.14

A canker is an inverted blister on the bark of the tree which, during some time of the year, may ooze sap. A canker forms on branches or trunks of trees. Fungi, very small organisms that produce spores and mold-like material, cause most canker diseases. Leaves of a tree affected by these fungi begin to turn yellow and may drop to the ground. Some limbs may not develop new leaves in the spring. In severe cases, trees may die if canker disease isn’t treated for a few years. Canker diseases can be controlled if diagnosed early. When leaves turn yellow prematurely, check the limbs and branches for damage or bark discoloration. These areas may be shrunken or shriveled. Check inside these areas for little bumps or pimples breaking through the bark tissue. They’re evidence that a fungus is causing the canker. To control canker disease on trees, cut off the affected branch or limb. If a large canker is on the main trunk, the tree may need to be replaced. No effective chemicals are available to control the fungi that cause canker disease. Good management practices, including fertilizer, proper drainage, watering and appropriate tree selection, are important for healthy trees and are the best defense against fungi. Cytospora Canker Cytospora canker is caused by various species of the fungus Cytospora (sexual genera of Valsa and Leucostoma). These pathogens affect many species of shrubs and trees in Colorado, including aspen, cottonwood, lombardy and other poplars, apple, cherry, peach, plum, birch, willow, honeylocust, mountain ash, silver maple, spruce, and Siberian elm. Some Cytospora species are host-specific while other species can infect several

different tree species. For example, willow, cottonwoods, and aspen are susceptible to one species. The fungus attacks trees or parts of trees that are injured or in a weak or stressed condition. The fungus grows in the living bark (phloem) and wood (xylem) and kills by girdling the branch or tree. The fungus can attack tree bark during the fall-winter spring seasons when temperatures are warm but the tree is dormant and cannot defend itself. Trees affected by drought, late spring frosts, insect and fungi defoliation, sunscald, herbicides, or mechanical injury are susceptible to Cytospora infection. The disease especially affects trees with root damage, which are often found in areas under construction, or trees that recently have been transplanted. Stands of aspen that have been thinned and young aspen sprout stands may suffer from Cytospora canker. Sexual and asexual spores of Cytospora species infect freshly wounded tissue. The spores are released after fruiting bodies have absorbed water during rain events. Conidia ooze out of the wet fruiting bodies and are dispersed by rain splash and blown by wind. Many times fruiting bodies are not formed since the cankered tissue dries out too rapidly in the dry western climates.

Fig. 1: Orange discoloration found with cytospora canker

Symptoms Cytospora species cause branch dieback and cankers on trees or shrubs. Cankers on stems and branches are often elongate, slightly sunken, discolored areas in the bark. Many times, however, the discoloration is not evident because the fungus killed the bark rapidly. The fungus grows so fast on stressed trees that there is no evidence of a sunken canker. Bark often splits along the canker margin as the tree is defending itself and callus formation occurs. The fungus may quickly girdle and kill twigs without

forming cankers. Symptoms vary with host species affected and stage of disease development. Bark above infected cambium may appear sunken and yellow, brown, reddish-brown, gray, or black. Diseased inner-bark and cambium turns reddish-brown to black, and becomes watery and odorous as it deteriorates. Wood below the cambium is stained brown. Liquid ooze on aspen and gummy ooze on peach and cherry are common. Cankers, sunken dead areas of bark with black pinhead-sized speckling or pimples, may be evident. The pimples are the reproductive structures of the fungus. Under moist conditions, masses of spores (seeds) may ooze out of the pimples in long, orange, coiled, thread-like spore tendrils. Reddish-brown discoloration of the wood and inner bark also may be evident. Dead bark may remain attached to the tree for several years, and then fall off in large pieces. On spruce trees, the disease appears as sunken, resinous areas surrounded by swollen callus, giving a gall-like appearance. Small black fruiting bodies may occur on the canker. Once the branch is girdled, needles may yellow or redden. The branch eventually dies. Large amounts of resin flow from infected areas, coating branches and stems. Unless you see sunken areas surrounded by swollen callus, resin flow on spruce may indicate that other stresses, diseases or insects are affecting the tree. Control Because this canker disease usually occurs on a weakened host, the primary method of control is to prevent stress on the tree. Drought and oxygen starvation of roots by flooding soil with water are the two most common stresses that predispose trees to Cytospora infection. High temperatures seem to be related to Cytospora canker on our local alders. To help a tree resist infection, prepare soil before planting, fertilize, water properly for winter and summer, prune, and avoid injury to the trunk and limbs. Proper care of recently transplanted trees also is essential to avoid stress and infection. Wounds caused by lawnmowers and weed trimmers are prime targets for infection on trees in landscaped areas. Insects, such as oystershell scale, stress the tree and predispose it to Cytospora infection. Insects should be controlled to prevent mortality by the combined stress of the insects and Cytospora canker.

Help prevent cankers at pruning wounds on peach and cherry trees by applying labeled fungicides as wound dressings. Do not rely on the effectiveness of fungicides on wounds of other trees to prevent infection. Another way to prevent Cytospora damage is to use species or varieties well adapted to the planting site conditions. These cultivars will be more likely able to resist the disease. Purchasing healthy nursery stock will decrease the possibility of infection. Once infection occurs, the best treatment is to increase plant vigor and sanitation. Remove all infected limbs and other areas. When removing branches, arborists and homeowners should make a smooth cut at the base of the limb, as near the trunk as possible, without damaging the branch collar (swollen area at base of branch). Jagged and rough cut surfaces promote infection. Once infection occurs, the best treatment is to increase plant vigor and sanitation. Remove all infected limbs and other areas. Clean wounds to avoid further spread of infection. Remove dead bark to dry out the diseased area and help the tree defend itself against insect and fungal attacks on the cankered area. Directions for proper wound and canker treatment are as follows: • Prune or cut trees only during dry weather. • Clean tools and wipe them with ethyl alcohol, Lysol or other disinfectant. Clorox may be used at a concentration of one part Clorox to nine parts water. • If a wound is fresh (one month old or less), use a sharp knife to carefully cut and remove all injured or diseased bark back to live, healthy tissue. If the wound is older, just remove loose bark pieces. It is important not to cut, remove or damage callus that may be forming at the canker edge. Callus will look like swollen bark growing across the dead area. Scrape the wound surface clean of loose bark. • Clean tools and disinfect after each cut. • Cleaned wounds should not have any sharp angles. • Do not apply any tar, oil-based paint or other wound dressing. The best method to prevent infection or decay is to allow the cleaned tissue to dry out. Fireblight Fireblight is a bacterial disease that affects apple, crabapple, mountain

ash and pear trees. It can kill branches and even whole plants if not treated. Springtime weather that is warm and rainy encourages fireblight. Bees and other pollinators may introduce the disease to blooming trees during pollination. As a result, symptoms often begin in the blossom, which develop brown, mushy-looking petals. More commonly, trees with fireblight develop curling, bending and blackening shoots, called shepherd’s crooking. Leaves turn yellow, then brown and finally black but remain on the branch. Selecting fireblight resistant varieties of apples, pears, and crabapples is a good way to minimize the chances of fireblight infection. Honeylocust Diseases Thyronectria and Black spot Nectria cankers are common on honeylocusts and occasionally Coral spot Nectria cankers are noted.

Fig. 5-16: : Asexual fruiting mats pushing through bark lenticels of Thyronectria canker

Thyronectria canker is caused by the austroamericana (Thyronectria austroamericana).

fungus

Pleonectria

Black spot Nectria canker is caused by Nectria nigrescens (Tubercularia ulmea). Coral spot Nectria canker is caused by Nectria cinnabarina. Root collar rot, a disease caused by soil microorganisms, kills the bark and outer wood on honeylocusts at the ground line, essentially girdling the tree by a canker. All ages and cultivars of honeylocust, including thornless and podless cultivars, are susceptible to cankers and collar rot. Symptoms and Signs

Cankers Disease symptoms include dieback of affected branches, reduced foliage, yellow foliage, premature fall coloration and early leaf drop. Cankers (areas of dead tissue) are found at the base of trees, at branch crotches, around wounds or on branch stubs. Cankers can range from slightly flattened surfaces to distinctly sunken areas with large callus ridges at the canker margin. Areas of stems and branches with thin bark may have a red-yellow discoloration. The condition of the bark and cambium (the tree’s growth tissue, between bark and wood) can indicate the presence of a canker. Infected bark and cambium will be loose and wood beneath them may have a dark, wine-red to yellow discoloration instead of a normal white or light color. The reddish color associated with the center of honeylocust stems is not related to these diseases. Diagnosis of the disease is easier if fruiting bodies (the spore-producing structures) of the fungi are present. Look for fruiting bodies in areas of the bark that have been dead for a year or less. Samples should be sent to a plant health clinic for culturing or microscopic confirmation. Thyronectria cankers caused by Pleonectria austroamericana have bumpy, cushion-like asexual fruiting bodies that are light yellow-brown when fresh but blacken with age. It also produces sexual fruiting bodies (perithecia) that are reddish-brown and also darken. Fruiting bodies usually are found in bark openings, such as lenticels (raised areas of bark that act as breathing pores) and scattered on bark surfaces in thinbarked areas. Large lenticels should not be confused with fruiting bodies. Black spot Nectria cankers, caused by Nectria nigrescens, appear very similar to Thyronectria cankers, and are best distinguished from P. austroamericana by cultural characteristics, morphology of the asexual fruiting structures, and ascospore morphology. The asexual fruiting bodies are raised bumps on the bark (sporodochia) and are creamy to peach colored when fresh, but normally turn dark brown to black within a few days of drying conditions. They usually form under a thin layer of bark or are exposed on the bark surface. The sexual fruiting bodies (perithecia) are round flask shaped and reddish brown, solitary or up to 20 on a raised mass of fungal tissue (stroma) and can be found clustered at base of sporodochia. Coral spot Nectria cankers caused by Nectria cinnabarina appear

similar to the two other cankers butN. cinnabarina cankers may contain raised smooth fungal masses (sporodochia) that are creamy to coral colored when young (usually for several weeks) and tan, brown or black when mature. The sexual fruiting bodies that look like small round flasks (perithecia) form in late summer singly or in groups up to 15 and are bright red to reddish-brown. Root Collar Rot Symptoms of root collar rot need to be recognized promptly because the disease can rapidly kill trees. Early fall coloration of a portion of the tree may indicate a large amount of damage. Small drops of gum on the stem near the ground or farther up the stem usually indicate that collar rot girdling occurred below that point. Loose bark and discolored wood (yellow to brown instead of white) just below the bark indicate initial collar rot and are the most indicative symptoms. Extensive death and discoloration of bark and wood can occur over several months. Black spot Nectria or Thyronectria cankers at the tree’s base usually indicate collar rot is active or was active in the past. Disease Cycle Cankers These fungi overwinter on infected trees as vegetative material (mycelium) and fruiting structures. Since the fungi also can live in dead tissue, they can become established or produce spores on dead wood such as branch stubs, wound edges or firewood. High humidity and wind-driven rain favor spore release and infection. Infections may take place through branch crotches, pruning wounds or other wounds in the bark. The fungus grows in the bark, cambium, and outer wood, where it eventually kills the cambium and surrounding cells. Death of the tree or affected parts occurs because of cambial death. Fruiting bodies can form within one month after the tree bark is killed and are abundant on dying or dead trees. Root Collar Rot Frequent watering in heavy clay soils may induce soil microorganisms to kill the bark and cambium at the tree base just below ground-line. Thyronectria or Black spot Nectria fungi may then infect the weakened tree above the area previously killed by collar rot. Damage and Control

Cankers at the tree base usually are fatal. Main stem or branch crotch cankers may completely girdle the tree, depending on the tree’s health. Stressed trees cannot stop the fungus, whereas healthy trees may be able to stop canker expansion and recover. Root collar rot is common in urban areas in Colorado and nearby states and is responsible for the death of many of the honeylocusts killed by disease. Cankers The best way to control cankers is to prevent wounds and promote tree vigor. Any injury to the base of a honeylocust is potentially an entry point for fungi. Lawnmowers, weed trimmers and construction work commonly cause basal injuries. Injuries to the stem and trunk, such as those caused by squirrel gnawing, pruning and sunscald, can be minimized by proper action. Should physical damage occur, remove loose bark and allow the wound to dry. A variety of stresses predispose honeylocusts to infection by canker fungi. To help prevent infection, avoid stress due to improper planting practices, drought, overwatering and insufficient area and oxygen for root growth. How about this. In general, planting small trees such as 2 to 9 cm diameter trees (1 to 3 inches) will ensure successful establishment rather than planting large trees >10 cm (>4 inches). Water trees adequately with about 2.5 cm (1 inch) of water per week. Avoid over watering trees since their roots need oxygen. Prune dead or infected branches to reduce the chance of other infections. Prune cankers on limbs by cutting at a branch junction and at least 30 cm (1 ft) below the visible margin of the canker. Prune in cool, dry weather to minimize reinfection. Cut out small cankers on main stems. Remove dead or dying bark and discolored wood. The area of bark removed should extend 2.5 cm (1 inch) into healthy tissue. If the tree appears to be recovering, however, do not cut into healthy tissue. Wound dressings are not recommended. Disinfect all tools used to prune and cut. Spray with Lysol® or dip in 70 percent rubbing alcohol or a 10 percent bleach solution (one part bleach to nine parts water) and dry after each cut. Prompt removal of all infected trees reduces the chance of spreading the infection. Because canker fungi can grow on dead wood and produce spores

that can infect nearby trees, keep the wood dry, bury it in a landfill or burn it within three weeks of cutting. Research shows Sunburst honeylocust is the most susceptible to cankers, while Imperial, Skyline, Trueshade, and Thornless are most resistant. Root Rots Root rots damage plants by stressing or killing root systems. Two common soil-inhabiting fungi that cause root rots include Fusarium sp. and Rhizoctonia sp. Root symptoms of these (and other soil-borne) fungi include darkening, limpness and mushiness. Rotted roots may break off easily. The cortex (the outer protective covering) of roots sloughs off, leaving behind the thread-like root core. Leaves, stems, and entire plants may wilt, prompting one to think that the plant simply needs more water. (Unfortunately additional water often makes the problem worse.) Generally, the lower, interior leaves turn yellow, then brown and drop off. In addition, plants may be stunted. If enough roots are damaged, the plant eventually dies. There are no root-rot resistant plants. Management strategies include avoiding overwatering and improving soil drainage. Roots stressed from overwatering or oxygen starvation easily succumb to root rots, since the organisms move through moist soil and water. Sometimes a plant with root rot may be salvaged by cutting off damaged roots and replanting in well-drained soil. Biological pesticides containing hyperparasites may help protect against root rot. These products are not designed to “rescue” plants from ongoing damage, but act as preventives. In the Green Industry, root rots can be managed with a combination of the cultural management strategies and through use of fungicides. Because not all fungicides kill all root rot fungi, it is essential to determine which root rot organism is causing the problem through microscopic examination, so the correct product can be recommended.

BACTERIAL DISEASES

Bacteria are single-celled microorganisms. They contain no nucleus and reproduce by dividing in two equal parts (fission). As a result, they multiply and mutate rapidly. Bacteria function as either parasites or saprophytes. Bacteria can infect all plant parts. Unlike fungi, bacterial must find a natural opening for entry. Bacterial cells can move from one plant to another in water, soil, and plant material, just as fungi do. However, bacterial pathogens are more dependent upon water. Conditions must be very wet and/or humid for them to cause significant and widespread damage. Bacteria move between plant cells and secrete substances that degrade plant cell walls so the contents can be utilized. Some produce enzymes that break down plant tissue, creating soft rots or water-soaking. Like the fungi, bacteria cause symptoms such as leaf blights and spots, galls, cankers, wilts and stem rots. Bacterial leaf spots appear different from fungal leaf spots due to their intercellular movement. Veins often limit the development of a lesion, so they appear angular or irregular, not round. Bacterial diseases are not common in the Rocky Mountain region due to lack of natural moisture. It is difficult for beginners to tell fungal and bacterial plant symptoms apart. Table 5.1 may be used to help distinguish symptoms caused by these pathogens. Table 5.1.Comparison of Fungal and Bacterial Leaf Spots Symptom Description

Fungal

Bacterial

Water-soaked appearance

no

yes

Texture

dry, papery

slimy, sticky

Smell

none

yes

Pattern

circular, target-like

irregular, angular

Disintegration

no

yes

Color change

red, yellow and purple halos

non

Structures of pathogen

mycelia, spores, fruiting structures

non

Common bacterial diseases include Bacterial Wetwood (Slime Flux), Fireblight (Erwinia amylovora), and Bacterial Leaf Spot (Erwinia sp.). For additional information, refer the following:

• Bacterial Wetwood • Fire Blight Bacterial Wetwood Bacterial wetwood, also known as slime flux, is a common disease that affects the central core or bark of many shade and forest trees. Wetwood is common on elm, cottonwood, aspen and willow, although it may also affect ash, fir, maple, apple and poplars. To identify wetwood, look for a yellow-brown discoloration on the trunk or branches. The area of infection is moist and under high internal gas pressure. This combination of high pressure and moisture causes slime to ooze from the tree’s central core where the problem originates. Fresh ooze is often rank, slimy and attractive to yeast fungi and insects. Dried ooze has an ashen-gray appearance. Wetwood slime is toxic to the tree’s cambium, the tissue in woody plants that produces new cells. This means the disease will inhibit new growth where the tree is infected. Wetwood slime also is lethal to foliage, young shoots and grass. Wetwood-infected tissue does not greatly alter the wood strength of most trees. The low-oxygen environment created by the bacteria deters woodrotting fungi that otherwise would weaken the tree. However, wetwood causes boards that have been cut from affected trees to warp and split when they’re dried. The infection process for wetwood is not well-understood. Although oozing often is associated with wounding or pruning, it’s unknown whether the tree is invaded upon injury or if the infectious microorganisms are already present. Bacteria associated with wetwood are common in soil and water, and probably enter trees through root wounds. The oozing could transfer bacteria, causing new stem or branch wounds. Wetwood also may occur in seedlings that develop from infected seeds. Although insects feed on the slime, they cannot transmit the disease. Unfortunately, no effective control measures exist. The best defense is to maintain tree health and avoid mechanical damage to stem and roots. Drought conditions may increase wetwood problems. Be sure to water and fertilize infected trees especially if the tree shows nutrient deficiencies.

A tree may survive several years with wetwood. However, because bacteria prevent new growth and inhibit the transport of water and nutrients, the tree eventually will succumb to the disease. Fire Blight Fire blight is a bacterial disease that affects certain species in the rose family (Rosaceae). It is especially destructive to apples (Malus spp.), pears (Pyrus spp.), and crabapples (Malus spp.). The disease also can occur on serviceberries (Amelanchier spp.), flowering quinces (Chaenolmelesspp.), cotoneasters (Cotoneaster spp.), hawthorns (Crataegus spp.), quinces (Cydonia spp.), pyracanthas (Pyracantha spp.), blackberries (Rubus spp.), raspberries (Rubus spp.), and mountain ashes (Sorbus spp.). Disease incidence varies from year to year and severity is influenced by cultivar susceptibility, tree age, succulence of tissues and spring meteorological conditions. The disease is most serious when spring temperatures during pre-bloom and bloom are warmer than average. Warm rainy springs are particularly conducive to rapid spread of the pathogen, resulting in blossom blight. Blight of twig terminals can occur in late May through June during wind driven rain events. Hail and wind damage provide wounds that allow the pathogen to enter at other times. Hot summer weather generally slows or stops the disease. Disease Cycle Fire blight is caused by the bacterium Erwinia amylovora. The bacteria overwinter in blighted branches and at the edge of cankers (areas of bark killed by bacteria) (Figure 5-17). In spring, when temperatures frequently reach 65 F, the bacteria multiply rapidly. Masses of bacteria are forced through cracks and bark pores to the bark surface, where they form a sweet, gummy exudate called bacterial ooze. Insects such as aphids, ants, bees, beetles, and flies, are attracted to this ooze, pick up the bacteria on their bodies, and inadvertently carry the bacteria to opening blossoms. Bacterial ooze splashed by rain can also spread the pathogen.

Fig 5.17: Fire blight life cycle. Once in the blossom, bacteria multiply rapidly in the nectar and eventually enter the flower tissue. From the flower, the bacteria move into the branch. When the bacteria invade and kill the cambial tissue of the branch, all flowers, leaves and fruit above the girdled area die. Infection also can take place through natural openings in leaves (stomata), branches (lenticels), pruning wounds, insect feeding and ovipositing, and hail. Droplets of bacterial ooze can form on twigs within three days after infection. Diagnosis Symptoms of fire blight are first seen about the time of petal fall. Infected blossoms appear water-soaked and wilt rapidly before turning dark brown; this phase of the disease is referred to as blossom blight. As the bacterial invasion progresses, leaves wilt, darken and remain attached to the

tree; this gives the tree a fire-scorched appearance, thus the name “fire blight.” Monitoring for Diseases Regular plant inspection, especially on lower and inner leaves, will alert gardeners to foliar problems. Foliar diseases are progressive- they begin as small spots on a few leaves. Lesions grow and coalesce and may cause leaves to yellow and die. Identify problems early on to determine the cause of the problem. Monitor affected plants through the season. As a result, it is necessary to find new methods of crop protection and disease combating which take the condition of the plant into account. Wageningen UR Greenhouse Horticulture researchers are working on alternative methods and applications based on prevention (business hygiene), antagonism and competition (other bacteria or fungi), climate control (ventilation), or physical solutions (light). Management: Many bacterial, fungal and viral diseases attack vegetable crops in Maryland home gardens. Most of these are not serious and in very few cases is spraying a fungicide recommended. Remember, when disease symptoms are noticed it is usually to late to spray a fungicide. Below are twelve tips that can help you prevent disease problems: 1.

Select disease-resistant varieties, particularly for those diseases that appear in your garden each year.

2. Purchase certified, disease-free potato tubers, garlic bulbs, and asparagus and rhubarb crowns. 3. Avoid planting on wet, poorly drained sites. 4. Pull soil up into raised beds if drainage is not very good. 5. Dig or till compost into the soil each year. 6.

Grow healthy plants by providing adequate light, water and nutrients. Give each plant adequate space to ensure good air circulation and add organic matter to your garden each year.

7.

Keep bare ground covered with an organic mulch. Newspaper covered with straw works very well.

8. Avoid watering foliage in the evening. It is best to direct irrigation water

around the plant base where it can quickly reach the root zone. 9. Avoid handling wet foliage. Harvest your vegetables before they become over-ripe. 10.

Cut off and discard leaves and pull up and discard entire plants that are badly infected by disease.

11. Pick-off and remove diseased fruits and clear your garden at the end of the season of all plant debris. This should be composted or tilled into the soil. Plant parts infected with especially damaging diseases, like late blight of tomato and potato, southern blight, and white rot, should be bagged and put out with your trash. 12. Keep weeds to a minimum and control those insect pests like thrips, aphids, flea beetles and cucumber beetles that are most likely to spread diseases. When disease symptoms are observed it is often too late to apply a fungicide, although fungicide treatments can help to protect new or uninfected foliage. Fixed copper, sulfur, and horticultural oil are some organic fungicides used by home gardeners. Always, carefully read and follow all pesticide label information and test the spray on a small part of the crop to check for signs of leaf injury (phytotoxicity.) Management of Biotic Plant Disease Plant disease is best managed through an integrated approach, which includes a combination of cultural, mechanical, biological, and chemical practices. •

Cultural management includes appropriate plant selection. Utilize plantsthat perform well in the local climate. Use disease resistant varieties whenpossible. Plant certified seed or seed pieces. Place plants in the appropriate environment for optimum growth. For example, grow shade-loving plants in the shade, not hot sun. Prepare soil before planting to improve root growth, reduce compaction in clay soils, and improve water holding of sandy soils. Apply fertilizer and water according to plant needs. Prune correctly, as needed, and at the correct time of year. •

Biological controls include the use of compost, compost teas, andhyperparasite products, which may reduce pathogens by

introducingbeneficial microbes. Encourage beneficial insects by planting floweringplants attractive to all stages of development. Avoid blanket applicationsof pesticides, which may kill beneficials in addition to harmful insects. Spottreat pest problems instead.

BIOLOGICAL FUNGICIDES Biological fungicides (“biofungicides”) are composed of beneficial microorganisms including specialized fungi, bacteria and actinobacteria (filamentous bacteria) that are used against fungi and bacteria that cause plant diseases. Many of these microorganisms are found naturally occurring in soils. Researchers have isolated specific strains which have been formulated with additives to enhance their performance and storage. More greenhouse growers are incorporating biological fungicides into their disease management programs. However, it is important to understand how biological fungicides differ from conventional fungicides, to understand their benefits and limitations in order for them to be an effective part of your disease management plan. Biological fungicides are living organisms that are best used preventively before disease occurs and not as a rescue treatment for diseased plants. They are best used in conjunction with good cultural practices, proper sanitation and promotion of plant health. Biofungicides Work There are a number of ways in which biofungicides work (their mode of action) including direct competition or exclusion, antibiosis, predation or parasitism, induced resistance and plant growth promotion. Many biological fungicides work in multiple ways, such as by competition and parasitism, so are less likely to develop resistance than conventional fungicides that often work in a single way with a specific mode of action. •

Direct Competition/ Exclusion before root infection can occur, pathogens must gain accesses to the zone closely associated with the root, known as the rhizosphere. For foliar diseases, the pathogen must make contact with the leaf or flower zone. The biofungicide grows a defensive barrier around this root, leaf or flower zone. The beneficial microbes competes with plant pathogens for nutrients, infection sites

•

•

•

•

and space, excluding the pathogen. Antibiosis the biofungicide produces chemical compounds or secondary metabolites such as antibiotics or other toxins that kills the target organism. The biofungicide create compounds that inhibit fungal or bacterial spores from germinating and causing plant disease, or the compounds restrict the pathogen’s growth. Predation or parasitism the biofungicide attacks and feeds on the pathogen, producing cell wall degrading enzymes, inhibiting or killing the pathogen. Induce Resistance to the host plant the biofungicide triggers the host plant to turn on its own defense mechanisms. Plants produce salicylic acid (a derivative of aspirin) which travels to other parts of the plant and signals these tissues to activate their natural defense mechanisms. This is also known as systemic acquired resistance (SAR) which improves the plants response to pathogen attack by priming the metabolism of plant defense compounds. Plant Growth Promotion - the biofungicide promotes root and shoot growth in the absence of disease causing pathogens. There may be increased nutrient availability of iron and other micronutrients by changing the pH or enzymes help dissolve insoluble elements.

Some Common Beneficial Microorganisms that are Commercially Available Beneficial fungi such as Trichoderma have been isolated from soil, decaying wood and plant organic matter. Different species are commercially available including T. harzianum and T. virens. Dormant spores of Trichoderma are applied, the spores germinate and the fungal mycelia coils around plant roots blocking the pathogen which results in a barrier to infection. The fungus also attacks the pathogen by secreting enzymes that attack cell wall of pathogen. There is also enhanced plant and root growth so the fungus has more roots to colonize. The combination of T. harzianum and T. virens has shown suppression of P. aphanidermatum, with more benefit against Phytophthora than T. harzianum alone. Gliocladium catenulaturm is a fungus isolated from Finnish field soil. It colonizes the leaf and root surface. Gliocladium works by hyperparasitism and competition for nutrients and space.

Bacteria Bacillus subtilis is a naturally occurring saprophytic bacterium with different strains commercially available. Bacillus subtilis works in a number of ways as it produces antibiotics, displaces the pathogen by inhibiting spore germination and interferes with the attachment of the pathogen to the plant. It also improves plant immunity and signals these tissues to activate their natural defense mechanisms or Induces SAR against bacterial pathogens. When combating bacterial diseases, growers can alternateBacillus with copper fungicides to help reduce the potential for plant damage or phytotoxicity that may occur from repeated sprays of some copper products under certain conditions. Bacillus can also be used against fungal leaf spots. Bacillus amylolquefaciens colonizes the plant rhizosphere, stimulates plant growth and suppresses competing fungal and bacterial pathogens. Streptomyces is a filamentous bacteria found in soil and decaying vegetation that produces spores and produces antibiotics. Streptomycin takes its name directly from Streptomyces. Streptomyces grisevirichis K 61 was originally isolated from sphagnum peat and S. lydicus WYEC 108 is a naturally occurring bacterium found in the soil. Benefits of Biological Fungicides • Reduced risks to applicators and the environment • Shorter re-entry intervals and days to harvest intervals than many conventional fungicides • Many are labeled for use on edible crops including herbs and vegetables • Less chance of plant damage, but not always, so consult product labels • Generally compatible with beneficial predators and parasites (natural enemies), beneficial nematodes (check company websites for more information) • Improved nutrient uptake of certain elements • Can be used in rotation with conventional chemicals to reduce the risk of pathogens developing resistance to conventional fungicides (especially systemic fungicides) • Not genetically modified Limitations of Biological Fungicides

• Must be used preventively, for they will not cure diseased plants • Must be used with proper cultural controls for plant growth, including clean starter material • Must use strict sanitation protocols • Have a shorter shelf life (consult labels) than conventional fungicides and need to be stored under proper conditions • May need to be re-applied more often than conventional fungicides • May need to integrate traditional fungicides in rotation for more aggressive pathogens such as Thielaviopsis andPhythopthora or stem rots such as Rhizoctonia or Phytopthora Biofungicides Start with clean greenhouse and clean starter material. Biological fungicides must be used as a preventive treatment in growing media or as a foliar application. For foliar diseases, it may be helpful to combine their use with the selection of disease resistant cultivars for disease suppression. Apply immediately after mixing with water. Check company websites for compatibility information with other materials. Because biofungicides are living organisms they have a limited shelf life and need to be stored under proper conditions. Do not stock pile biofungicides and be aware of the expiration date on the package. In university studies, researchers sometimes see an uneven effect when applying biological fungicides, however these studies are conducted with higher disease pressures than in commercial greenhouses. In order to complete your own in-house trials, leave a number of plants untreated to serve as your control treatment. Differences in your crop, potting mix, media pH, fertilizer use and disease pressure may influence how well these different products work for you. Use in alternation with conventional fungicides. Biological fungicides are a useful tool for growers provided that they are used preventively, in combination with proper sanitation and good cultural practices. In the future, more combination products may become available.

6: Diseases of Greenhouse Ornamental Crops AERIAL BLIGHT (Phytophthora parasitica) Most susceptible plants: vinca. This fungal pathogen causes a major disease in vinca in the landscape but it can also be a serious disease of greenhouse-grown vinca. Symptoms

Phytophthora parasitica

Initial symptoms of infection occur on leaves. Leaf infection is characterized by a rapid collapse of the leaf. Infection progresses to the leaf petiole and to the location of the attachment of the petiole to the plant stem. A brown, sunken stem lesion develops at the point where the petiole attaches to the stem. This brown stem lesion develops on the stem causing stem collapse. If wet conditions persist following plant infection, the fungus will grow to the base of the plant resulting in plant death. A unique feature of Phytophthora

aerial blight is its decided aerial nature; this fungus rarely causes a primary root rot but causes massive damage to the aerial portion of the plant. Control Control of this disease has been very difficult and has proven impossible in many situations. Most effective control in the greenhouse will require that the plants are grown on benches, as far above soil as feasible. Remove symptomatic plants as soon as they become obvious on the greenhouse bench. Although foliar-fungicides have not been totally satisfactory, and have failed totally in the landscape for control of this disease, application may be useful in greenhouse situations if applied at the first sign of disease.

BACTERIAL BLIGHT (Xanthomonas campestris) Most susceptible plants: begonia, geranium, zinnia. This disease can be devastating, causing loss of an entire crop. The pathogen is systemic so watch for symptoms and take immediate action upon detection. Symptoms

Xanthomonas campestris

Leaf spots begin as water-soaked blotches on leaves, frequently with large areas of chlorosis and browning at the leaf margins. An inverted “V” pattern commonly develops soon after initial symptoms occur; the “V”shaped lesion usually turns a copper color as the infection proceeds. Leaf collapse is common as is the development of a soft, watery deterioration of the plant stem. Plant wilt and decline follows.

Control The most important management strategy for this disease is to avoid introducing the pathogen into the greenhouse. Use disease-free, cultureindexed cuttings. Know the typical symptom to look for. There is no effective chemical control. Aggressive roguing of infected or symptomatic plants is important. Keep foliage dry; avoid overhead wetting/irrigation.

BACTERIAL STEM ROT (Erwinia carotovora pv carotovora) Most susceptible plants: kalanchoe. Symptoms Infected plants develop a black, soil-line lesion that usually results in stem weakening, lower stem breakage, plant stunting, plant wilt and plant death. The soil-line lesion is usually very soft and mushy in texture.

Erwinia carotovora

Take care to avoid wounding the plant stem during transplanting or when spacing/moving pots. Chemical controls have not proven effective.

BLACK ROOT ROT (Thielaviopsis basicola) Most susceptible plants: pansy, vinca. Black root rot can cause significant production losses in greenhouse crops. Although this fungal pathogen also has a very wide host range, the most serious problems occurr on pansy and vinca. Pansy and vinca plug

infection has resulted in significant plant losses.

Thielaviopsis basicola

Symptoms The black root rot fungus damages the root of the plant, effectively interfering with the root’s ability to absorb nutrients. As a result of root injury, plants usually develop symptoms indicative of nutritional stress. Yellowing of the younger growth is a common symptom. Root examination of infected plants usually reveals a lack of healthy, white roots; infected roots are usually off-white, gray or black, depending on the stage and severity of infection. Control Control of black root rot can be difficult if the pathogen becomes established within the growing area. Pay strict attention to sanitation. Do not reuse plug trays or plastic pots. Store media in a location that is protected from contamination. Spot-check all plugs introduced into the growing area by carefully examining roots for healthy, white color. Stress has been shown to greatly enhance black root rot. Adverse temperatures, excessive moisture in the root zone, excessive levels of soluble salts and excessive use of fungicides or other plant production chemicals have all been implicated in intensification of black root rot. Because plugs are vulnerable to a number of stresses, all plugs should be planted as soon as possible after arrival. Several fungicides have proven effective. Use preventatively or at the first sign of infection for effective control. An acidic pH helps to manage black root rot; a pH range of 5.5-5.8 can reduce black root rot development.

BOTRYTIS BLIGHT (Botrytis cinerea) Most susceptible plants: exacum, geranium, impatiens. Probably the most common and troubling greenhouse pathogen is the gray mold fungus, Botrytis cinerea. Botrytis can infect any above-ground portion of the plant. Wounded or stressed tissues are much more susceptible to infection. Botrytis can cause serious problems in geranium both as a flower blight as well as a stem/cutting rot.

Botrytis cinerea

Botrytis blight, also know as gray mold, is a fungal disease caused by several species in the genus Botrytis. It affects the buds, flowers, leaves, and bulbs of many plants including: African violet, begonia, chrysanthemum, cyclamen, dahlia, geranium, lily, peony, rose, and tulip. The extent and severity depends on weather conditions and cultural practices. This disease is the primary cause of decay in cut flowers. Symptoms Diagnosis is usually straightforward and is characterized by the obvious gray-to-brown fuzzy growth on infected plant tissue. This disease is much more common when relative humidity is high, when air stagnation is present, when temperatures are cool and where free moisture occurs on plant parts. Control • Avoid injury or stress to plants. • Encourage good air circulation and air movement within the greenhouse. • Space plants so as to encourage air movement around individual plants.

• Use night heat plus ventilation at night to lower relative humidity within the greenhouse. • Keep temperatures above 60°C if possible. • Thorough greenhouse sanitation is mandatory! Remove infected plant material and plant debris. • Fungicide application may be effective if started early and if fungicides are rotated.

CROWN AND ROOT ROT (Phytophthora spp.) Most susceptible plants: gerbera daisy, gloxinia, pansy This disease is second only to Impatiens Necrotic Spot virus as a serious pathogen in gloxinia. Root and crown rot can also cause serious losses in gerbera daisy production. Infection can occur at any stage of gerbera production but seems to be more common after flowering begins.

Phytophthora spp.

Caused by soil-borne, fungus-like organisms (Phytophthora), crown and root rot occurs worldwide on almost all fruit trees as well as many woody ornamentals. Visible symptoms include: slow growth, sparse, yellowing foliage, small fruit, wilting in hot weather, or sudden plant death. The disease can be confirmed by using a knife to expose the inner bark of the root collar or large roots. Look for distinctive brown tissue (infected) in contrast to cream-colored tissue (healthy). The causal pathogen is present in most soils,

but only causes infection under optimal circumstances—high soil moisture or standing water, and susceptible host tissue. Once trees are infected, there is no cure. Symptoms Plants fail to grow adequately and usually remain noticeably stunted. The foliage develops an obvious off-color. Petioles may become infected where they attach in the crown area; as the petiole collapses, the attached leaf dies also. As the disease progresses, the entire plant wilts and dies. Phytophthora root and crown rot more commonly results in a rapid plant wilt, in which the plants appear normal and healthy one day but develop symptoms of rapid wilt and decline. The pattern in the flat or on the greenhouse bench is usually random but can be extensive, involving a large number of plants, especially if plants are not grown on benches. Overhead watering readily splashes the pathogen from plant-to-plant. Examination of the symptomatic plant will usually reveal a very wet root ball with significant root and crown discoloration and deterioration. Control Avoid growing plants on the ground or on mesh on the ground. Make sure that transplants are not planted too deeply: at the approximate same depth as the original plug not below the crown area. Avoid compaction of the rooting medium. Irrigate carefully; avoid overwatering. Drench-applied fungicides can be effective in managing this disease but should not be relied on to overcome poor cultural conditions.

FUSARIUM STEM AND ROOT ROT (Fusarium solani/Nectria spp.) Most susceptible plants: exacum Fusarium stem rot is caused by Fusarium solani. This is not a systemic disease, but a canker problem. Prevention would include avoiding deep planting or over fertilization, and m aintaining adequate soil calcium levels and pH 6.2 or above. Ammonium nitrogen sources will favor Fusarium.

Fusarium Stem Rot

Infection by this fungal pathogen can cause rapid wilt and plant decline. This disease can easily be confused with Impatiens Necrotic Spot Virus and Botrytis stem infection. Symptoms Typical symptoms develop as off-color foliage initially, with a progressive desiccation and wilt of branch sections or commonly, one side of the plant. Eventually, the entire plant may yellow, wilt and die. Examination of the lower stem commonly shows a whitish fungal growth in association with an obvious lower stem rot. Small, reddish round balls, which represent the Nectria stage (the sexual stage) of the fungus are commonly observed. Control Remove infected plants as soon as symptoms are observed. Preventative drench-applied fungicides have shown good control of this disease.

FUSARIUM WILT (Fusarium oxysporum f.sp. cyclaminis) Most susceptible plants: cyclamen Infection can occur at the seedling stage, with symptoms becoming obvious in older plants following an environmental trigger such as hot greenhouse temperatures. Symptoms

Yellowing of lower leaves

Infection is usually characterized by chlorosis of the leaf tissue, collapse of the leaf petiole, plant wilt and plant death. Lower leaves usually wilt and yellow first, followed by the rest of the foliage. The corm usually remains firm. The most distinguishing diagnostic feature of infection is the obvious brown-black discoloration of the internal tissue of the corm that can be seen if an infected corm is split lengthwise. Control This disease is best controlled by strict sanitation. Remove infected plants as soon as symptoms develop. Fungicides have not proven effective.

IMPATIENS NECROTIC SPOT VIRUS (INSV) Most susceptible plants: begonia, chrysanthemum, exacum, gloxinia, impatiens, vinca. This is one of the most serious diseases of greenhouse crops. INSV has a very large host range, numbering more than 648 different plant species. It is the number one disease of gloxinia and impatiens. INSV is transmitted by the thrips insect; it is not known to be routinely transmitted by any other means. One of the most frustrating features of INSV is that infection can result in a number of different symptoms. Symptoms

Symptoms of INSV infection may include black ring spots (impatiens), black foliar lesions (impatiens, vinca, cineraria), chlorotic ringspots (exacum, gloxinia, cyclamen), veinal necrosis (gloxinia, Aphelandra, impatiens), stem lesions (chrysanthemum, exacum), distortion of young growth, stunting and plant wilt. These symptoms can vary with the stage of growth of the host and with a variety of cultural conditions. Plants infected with INSV are systemically infected for the life of the plant. Although the plants are systemically infected, INSV can “compartmentalize” within its host, causing symptoms to occur only in a portion of the plant. Control INSV plants must be rapidly and thoroughly rogued from the production area to reduce infection to other susceptible plants. Monitor thrips activity routinely throughout the production area by the use of yellow or blue “sticky cards.” Destroy all weeds, both inside and outside the greenhouse, as weeds can serve as both a reservoir of the INSV virus as well as a habitat for the thrips vector. Control thrips activity by appropriate management strategies, including insecticides when needed.

POWDERY MILDEW (Erysiphe cichoraceaum) Most susceptible plants: begonia, chrysanthemum, gerbera daisy, kalanchoe, zinnia. Although most greenhouse crops can be infected by powdery mildew pathogens, each powdery mildew pathogen is specific to its host. The powdery mildew that infects gerbera daisy will not infect zinnia, etc. Powdery mildew is the most common disease of gerbera. Infection may start on the lower foliage and escape early detection if plants are not periodically monitored for this disease. Powdery mildew pathogens are readily disseminated in the air by air currents.

Erysiphe cichoraceaum

Conditions that favor the development of powdery mildew diseases are moderate temperatures with high humidity. Some powdery mildew pathogens are enhanced by fluctuations between warm and cool temperatures but a relative humidity of 85% is generally needed for disease to develop. Symptoms Powdery mildew diseases are very common to a number of greenhouse crops and are easy to diagnose by the development of anobvious white powder on the plant surface. An atypical brownish scab-like symptom also may occur. Control Fungicides can be effective in managing powdery mildew diseases but take care to initiate applications at the first sign of infection and to rotate among effective fungicides. Include a systemic fungicide in the spray rotation.

PYTHIUM BLACK LEG (Pythium spp.) Most susceptible plants: geranium Black leg is a stem infection of Geraniums (Pelargonium spp.) that results in a distinctive black discoloration of stems. As infected stems rot, they become soft and often bend over. This disease is caused by several species of the water mold Pythium. Pythium spp. are soil dwelling organisms that thrive in wet conditions and can survive in infected plant debris and soil. Black leg symptoms often start at the soil line and move up the plant.

Symptoms Pythium black leg develops as a distinctive blackened deterioration of the lower stem, starting at the soil line. Infected stem tissue softens and deteriorates, damaging the plant’s vascular tissue and interfering with movement of moisture to the leaves and other above-ground tissue. Plant wilt, stem collapse and plant death commonly result.

Pythium spp.

Control Take special care during transplanting to avoid excessive planting depth and wounding of stem and root tissue. Avoid excessive soil compaction during the transplant operation. Water carefully; avoid overwatering. Drenchapplied fungicides can be effective in controlling this disease problem.

7: Management of Fungus Gnats Fungus gnats (Bradysia spp.) commonly develop in the moist environments common in the greenhouse, especially in propagation houses and in newly planted plant material. Fungus gnats, Bradysia spp., were initially considered minor insect pests, primarily found in house plants potted in growing medium. They were not considered a problem in ornamental cropping systems. Today fungus gnats are recognized as major insect pests in greenhouses and nurseries and are one of the few insect pests in which the damaging life stage - the larva in this case - is located within the growing medium. They are especially a problem under excessively moist conditions during propagation, when plant cuttings or plugs are initiating root systems. Adults cause minimal plant damage, but females lay eggs that hatch into larvae that damage plants by root feeding. Both adults and larvae may spread plant pathogens.

BIOLOGY AND DAMAGE

Adult Fungus gnats

The fungus gnat life cycle consists of anegg, four larval instars, a pupa, and an adult.A generation may be completed in 20 to 28days depending on temperature. Adults arewinged, approximately 3 to 4 mm (0.011 to0.015

inches) in length, with long legs andantennae. Adults tend to congregate near thegrowing medium surface and live from 7 to 10days. Females deposit 100 to 200 eggs into thecracks and crevices of the growing medium.Eggs hatch into white, translucent, leglesslarvae that are approximately 6.0 mm (0.023inch) long with a distinct black head capsule.

Fungus gnat life cycle

The larvae are located within the top 2.5 to 5.0 cm (1 to 2 inches) of the growing medium surface or inside plant tissue. Fungus gnat larvae feed on plant roots, primarily the root hairs, and organic matter in the upper 2.0 cm (0.78 inches) of the growing medium. They may be distributed throughout the growing medium profile, even at the bottom of containers near drainage holes. Fungus gnat larvae also may emerge from the growing medium to feed on leaves lying on the growing medium surface or tunnel into plant crowns. The larvae prefer growing media with a “high” moisture content and require fungi as a supplemental food source in order to complete development. The type of food determines abundance and fitness of adults, and the reproductive capacity of females. Fungus gnat larvae feed on a wide-range of ornamental plants grown in

both greenhouses and nurseries, including Capsicum spp., Cyclamen spp., poinsettia (Euphorbia pulcherrima), Geranium spp., transvaal daisy (Gerbera jamesonii), Gloxinia spp., Impatiens spp., bedding plants, and vegetable transplants. Young plants and/ or seedlings are especially susceptible to injury from larval feeding, more so than mature plants, unless fungus gnat larval populations are extremely abundant. Larval feeding directly damages developing root systems and interferes with the ability of plants to absorb water and nutrients, resulting in stunted growth. Larvae also may cause indirect damage during feeding by creating wounds that allow entry of soilborne plant-pathogens. Additionally, both larvae and adults can transmit fungal diseases, including Botrytis spp., Pythium spp., Fusarium spp., Verticillium spp., and Thielaviopsis basicola from infected to noninfected plants. Fungus gnat adults may carry the spores of certain foliar and soilborne plantpathogens on their bodies, including B. cinerea, Fusarium oxysporum f. sp. radicislycopersici, T. basicola, V. alboatrum, and F. avenaceum. Adults can then disperse spores throughout a greenhouse or nursery. Fungus gnat larvae have been shown to ingest the propagules of Pythium aphanidermatum and macroconidia of F. avenaceum, which they disseminate or introduce into young healthy plants during feeding. It also has been reported that the oospores of Pythium spp. are able to survive passage through the digestive tract of fungus gnat larvae and are intact and viable (able to germinate) after being excreted.

DAMAGE Fungus gnats and shore flies are attracted to damp locations where fungi are apt to flourish. Fungi are a major part of their diet. Studies have shown that fungus gnats develop more rapidly and have greater survival on fungal diets. In the absence of a fungal food source however, fungus gnats are capable of feeding on healthy plant tissue. Fungus gnats are general feeders and can injure a number of flower crops grown in the greenhouse. Adults are primarily a nuisance however, larvae feed on plant roots, fungi and decaying organic matter and tunnel into the crown and stems of plants. The feeding damage creates wounds that allow soilborne pathogens to enter and can kill plants. Fungus gnat larvae may also carry some soil-borne pathogens such as

Pythium, Thielaviopsis andFusarium. Fungus gnats are a common problem on greenhouse crops gowing in media that contains a high percentage of peat moss or compost. Larvae present in infested plants or soil can lead to prolonged emergence of adults. Shore flies are not known to feed on healthy plant tissue. Adult and larval stages of shore flies feed primarily on algae or decaying organic matter and breed in moist environments.

IDENTIFICATION Adult fungus gnats are small (1/8 inch long), mosquito-like insects, with long legs and antennae. Their two wings are delicate and clear with a Yshaped vein in the wing pattern. Adults are weak flyers and tend to fly in a zig-zag pattern. They may be observed resting on the growing media surface or moving across lower leaves. Adult females are attracted to fungi so might be observed near plants with Botrytis sporulation. Females lay their eggs nearby so the developing larvae have access to a fungal food source. Fungus gnat larvae are small, (approximately ¼ of an inch long when mature), translucent to white in color with a distinctive black head capsule. Both fungus gnats and shore flies are common in the greenhouse. However, it is important to distinguish between the two, because management strategies differ. Shore fly adults (approximately 1/8 of an inch long), resemble a small housefly with stockier bodies, plus shorter legs and antennae than fungus gnats. Shore flies also have five distinct white spots which fungus gnats do not have. Shore fly larvae are white, wedge-shaped and do not have a distinctive head capsule. Larvae may be found near algae, a primary food source. They do not feed on plants.

FUNGUS GNAT DAMAGE Fungus gnat larvae feed on fungi and decaying organic matter, but so feed upon plant roots. This larval feeding is most damaging to seedlings, and young plants. Larvae also feed on the developing callus of direct stuck cuttings, delaying rooting. Fungus gnat larvae are general feeders. Plants with succulent stems, such as begonias, geraniums, sedum, coleus and poinsettias, are especially prone to injury and can suffer serious losses. As the young

feeder roots and stems are damaged, plants wilt and leaves turn yellow and drop. In laboratory studies, adult fungus gnats carried spores of Botrytis, Verticillium, Fusarium and Thielaviopsis as they moved from plant to plant. Spores have also been found in their droppings. It is unclear how important this disease transmission is in commercial greenhouses.

SCOUTING Monitoring is especially crucial if you are planning on targeting biological controls or insect growth regulators against the fungus gnat larvae. Inspect incoming plugs for fungus gnat larvae or their damage. Place yellow sticky cards in samples of growing media to monitor for any emerged adults. Yellow sticky cards, placed horizontally at the soil surface, can be used to detect fungus gnat adults. Check and change the cards weekly to detect early fungus gnat infestations. Use potato plugs (at least one inch in diameter) placed on the soil surface to monitor for fungus gnat larvae. When using potato plugs, place the plug so there is contact with the media to ensure that the potato plug does not dry out. To look for larvae, first check the growing media under the plug and then the surface of the potato itself. Check the potato plugs after 48 hours for the presence of larvae. Be sure to mark the locations where you placed the potato plugs, so you can easily find them! If not removed, potato chunks can “melt out,” sprout or be fed upon by mice. For smaller cuttings or plugs, potato slices, resembling a “French fry” can be placed in the growing media.

CULTURAL CONTROLS Adults are attracted to newly planted crops, making it important to thoroughly clean the greenhouse before introducing new crops. Dry, level, weed-free, well-drained floors help eliminate breeding areas. Keeping compost piles away from the greenhouse and cleaning up any spilled media on the floor also helps eliminate breeding areas. Avoid overwatering and allowing excess moisture to accumulate underneath greenhouse benches. Remove plant debris, weeds, and old growing media from inside and outside the greenhouse. Inspect incoming plugs for fungus gnat larvae or their feeding damage.

Recent studies have shown that fungus gnats may be introduced into a greenhouse from soilless media or rooted plant plugs. Adults are attracted to mixes with high microbial activity, or with high amounts of peat moss or hardwood bark. Avoid using mixes with immature composts less than one year old. However, no potting mix is completely immune to fungus gnat infestations. Adult females prefer to lay their eggs in protected, humid crevices in the media. How the media is handled and stored may be more important than the type of growing media used. If the growing media is stored outside and stays moist, it may support more fungus gnat activity. Tears or openings in the bags enable resident, native fungus gnat populations to gain entry into the media bags. Store the media so that it stays dry. Covering the growing media with a layer of coarse sand or diatomaceous earth does not help prevent egg laying by the adult females. Diatomaceous earth absorbs moisture from the growing media so that cracks develop where larvae can pupate and females can lay eggs. Biological Control Commercially available natural enemies include the soil dwelling predatory mite, Hypoaspis miles, (= Stratiolaelaps scimitus), the entomopathogenic nematode, Steinernema feltiae, and the rove beetle, Atheta coriaria. All should be used preventively and applied to moist growing media. Steinernema feltiae are beneficial, insect killing nematodes that are applied as a drench treatment against fungus gnat larvae. After entering the target insect through various openings, the nematodes multiply within the host and release a bacterium whose toxin kills the larvae. These beneficial nematodes reproduce within the fungus gnat larvae; exit the dead body and search for new hosts to infect. Fungus gnat larvae are killed in one to two days. A small, soil-dwelling predatory mite, Hypoaspis miles, feeds on fungus gnat larvae as well as thrips pupae and shore fly larvae. It is shipped in a vermiculite/ peat carrier with all stages of the predatory mites. The vermiculite/peat carrier can be distributed over the media surface, especially when pots are placed close together. These predatory mites are best used

when fungus gnat populations are low. The rove beetle is a generalist predator that feeds upon fungus gnat and shore fly larvae in the growing media. Adults are slender, dark brown or black and covering with hairs and have very short wing covers. Adults are nocturnal so are best released in the evening. Both adults and larvae tend to hide in cracks and crevices of the growing media. Bacillus thuringiensis var. israelensis, sold under the trade name of Gnatrol WDG, is most effective against the young first instar larvae. The bacteria must be ingested by the larva, after which a toxic protein crystal is released into the insect’s gut. Larvae stop feeding and die. Gnatrol WDG is only toxic to larvae for two days. Repeat applications, i.e. two or three applications at high rates, may be needed to provide effective control.

BENEFICIAL NEMATODES: EASY WAY FOR BIOLOGICAL CONTROL IN THE GREENHOUSE Growers that are interested in using biological control can start by using beneficial nematodes to manage fungus gnats. They are relatively easy to use and are applied in a similar manner to conventional pesticides with some special precautions that are listed in this factsheet. Nematodes are small, colorless, cylindrical round worms that occur naturally in soils throughout the world. Different species work best against different insect pests. Steinernema feltiae is primarily used against fungus gnat larvae and thrips pupae dwelling in the soil media. Fungus gnat larvae may be parasitized in any larval stage. Nematodes have traditionally been used against soil dwelling pests because they are sensitive to ultra violet light and desiccation. The beneficial nematodes enter the insect host through body openings. These insect killing nematodes multiply within the host and release a symbiotic bacterium (Xenorhabdus spp.) whose toxin kills the target pest, i.e. fungus gnats. The fungus gnat larvae are killed in one to two days by blood poisoning. More than one generation of nematodes may develop in dead host insect in the growing media. The infective juveniles then exit the dead body

and search for new hosts to infect. Use of Beneficial Nematodes The beneficial insect killing nematode S. feltiae is sold under the trade names of NemaShield, Nemasys, Scanmask and Entonem. All of these products are labeled as a soil drench treatment against fungus gnat larvae. Preventative applications to moist soils work best. •

Apply nematodes with a sprayer (remove screens and filters), injector, hose end sprayer or even a watering can in very small operations. • If using an injector, set the dilution to 1:100. Remove all filters or screens (50 mesh or finer) in any spray lines so that the nematodes can pass through unimpeded and undamaged. • If using a sprayer, keep spray pressure below 300 psi. • Although nematodes are applied in water, they are not aquatic animals and therefore they need extra care while in stock and tank solutions. Adequate aeration of the nematode suspension during application is important. This can be done using a small battery powered submersible pump or even mechanically with a stirrer to keep the solution agitated. A small pump will also keep them from settling on the bottom of the stock solution container, which they tend to do. The suspension in the spray tank should be kept cool and applied as soon as possible after mixing. This is especially important during the warmer months. The longer they are kept before spraying and the warmer the tank water, the more quickly their energy reserves are used up. Weaker nematodes are less robust during and after application, and less able to search for and infect a susceptible host. The symbiotic bacteria break down the host insect’s cuticle. The infected fungus gnat larvae rapidly disappear, so they may be difficult to locate in the growing media. Infected fungus gnat larvae are often opaque-white to light yellow in color. Use potato disks to monitor for fungus gnat larvae. Place disks on the surface of the growing medium two days before application in order to determine the population level prior to treatment, and again 3-5 days and 1012 days after application. Leave the potato disks on the growing media for two days in each case, before examining them for fungus gnat larval activity.

Beneficial Nematode use against western flower thrips Nemasys is also labeled for use against western flower thrips. In the late 1990s in the U.K., it was reported that cut chrysanthemum growers who applied nematodes weekly as a foliar spray, noted a reduction in their thrips populations. More recent work showed that soil-dwelling stages of thrips (especially the pupal stages) were highly susceptible to several species of nematodes, such as Steinernema feltiae. During the weekly sprays, a significant number of nematodes reached the growing media via runoff from the foliar sprays. Nematodes are very short lived on the foliage (significant reduction after one hour) but may persist for several weeks in the media. Mobile life stages on the plant (adults and larvae) appear to be less susceptible to attack. Thrips control noted in commercial crops may have occurred as a result of overspray and run-off into the soil after spraying. Special precautions are taken to help reduce potential desiccation: use of a non-ionic wetting agent such as Capsil, spraying in the late afternoon or evening, and the use of black shade cloth. Begin treatments with an early drench to the media and then continue with weekly sprays to the foliage making sure you have uniform coverage of the foliage and also sprench the soil. Specific tips for use against western flower thrips (from the Nemasys label) • Nematodes require moist conditions to enhance effectiveness. • If plants are dry, provide light overhead irrigation prior to nematode application. • Ensure good foliar coverage of spray mix to enhance contact with the target pest. • Use of a wetting agent or surfactant such as Capsil will enhance wettability of the spray mix and encourage nematode movement. • Following application, ensure that the crop remains wet for at least two hours. Note: Do not apply in direct sunlight. • Note: the nematodes will desiccate after about one day, depending upon environmental conditions. Grower feedback has been variable, with some observing excellent results and others less so. Efficacy will be variable depending upon the

relative humidity, and temperature in your greenhouse, dose applied, frequency of application, and life stage of the thrips. Some growers apply the nematodes with additional water in the summer months to ensure that the foliage stays wet to contact the thrips on the foliage. Depending upon the temperature, relative humidity levels and other environmental conditions, up to 2x the amount of water may be needed to keep the foliage wet for two hours. Regular monitoring, sanitation, proper spacing and judicious use of fungicides and biological fungicides may be needed to discourage foliar diseases. • Applying the nematodes as a heavy surface spray or “sprench” to young, incoming plant material will have an added benefit of targeting any incoming fungus gnats in the media as well as thrips pupae. • Growers who have had success with this application method, apply the nematodes on a weekly basis, and target the young growing point where thrips tend to hide. • As with any biological control measure, they are most effectively used preventively in conjunction with good cultural practices for thrips control (sanitation, rigorous weed controls, inspection of incoming plants and regular monitoring).

SUMMARY Fungus gnats (Bradysia spp.) are a major insect pest in greenhouse and nursery cropping systems because both the adults and larvae may cause direct and/or indirect plant damage resulting in possible economic losses. The challenge in dealing with fungus gnats is to approach management “holistically” by using strategies including scouting and growing medium selection, which are extremely important in alleviating problems with fungus gnats. Greenhouse producers and nursery managers who implement proper water management and sanitation practices - such as eliminating weeds and algae from production areas and properly timing the use of both insecticides and natural enemies - will experience fewer problems with fungus gnats in greenhouses and nurseries.

REFERENCES

Cloyd, R. 2015. Ecology of Fungus Gnats (Bradysia spp.) in Greenhouse Production Systems Associated with Disease-Interactions and Alternative Management Strategies. Insects. 6: 325-332. Ferguson, G., G. Murphy, and L. Shipp. 2014. Fungus Gnats and Shoreflies in Greenhouse Crops. Ontario Ministry of Agriculture and Food Fact Sheet 14-003. Lamb, E., B. Eshenaur, N. Mattson, and J. Sanderson. 2013. Practical Suggestions for Managing Fungus Gnats in the Greenhouse. Cornell University Ornamental IPM Factsheet. Meers, T. and R. Cloyd. 2005. Egg-Laying Preference of Female Fungus Gnat Bradysia sp. nr. coprophila (Diptera: Sciaridae) on Three Different Soilless Substrates. Journal of Economic Entomology. 98(6): 1937-1942.

8: Managing Aphids in the Greenhouse INTRODUCTION Aphids can be serious and persistent pests in the greenhouse. They are difficult to control due to their high reproductive capability and resistance to many different insecticides. Aphids are sucking insects that can cause curling and distortion of tender young growth. The presence of aphids, their white shed skins and honeydew can reduce the aesthetic quality of a wide range of greenhouse crops.

IDENTIFICATION Aphids are small (less than 1/8 of an inch long), soft-bodied, pear-shaped insects with long legs and antennae. Cornicles, “tail pipe like” protrusions, can be seen at the rear of their abdomen. Some of the most common species found in greenhouses include the green peach aphid (Myzus persicae), the melon or cotton aphid (Aphis gossypii) and the foxglove aphid (Aulacorthum solani). Potato aphids (Macrosiphum euphorbiae) may occasionally occur. Other species that growers may encounter include the gray cabbage aphid, the bright yellow-orange oleander aphid, and the reddish-brown chrysanthemum aphid. Tulip bulb aphids can infect many different bulbs in storage. Some aphids may even be found on plant roots (Pemphigus species). Proper identification is important in order to choose the most effective management option. Aphids vary in color depending upon the plants they are feeding on, so should not be relied upon to identify species. Green peach aphids have red eyes and may vary in color from pale yellow to green to pinkish-red. The pear-shaped adults are approximately 1/14 of an inch long. They have long cornicles that are approximately the length of their body and only slightly darkened at their tip. Green peach

aphids also have a pronounced indentation between the base of their antennae.

Melon or cotton aphids are generally smaller (less than 1/16 of an inch long) than green peach aphids. There is more variation in color within the same aphid colony. Melon aphids may be yellow to green to purplish-gray to black with distinctive white patches on the abdomen. Their short (approximately 1/3 of an inch long) cornicles are completely black. Melon aphids have antennae that are shorter than their body. Melon aphids do not have a distinct indentation are the base of their antennae like the green peach aphid. Growers frequently refer to melon aphids as “black aphids.” Foxglove aphids are also known as the glasshouse potato aphid. The pale green, shiny foxglove aphids have large dark green spots at the base of their cornicles. They also have black markings on their leg joints and antennae. Foxglove aphids also have an indentation between their antennae. They are larger in size than the green peach aphids (.11 of an inch) long. They are pale green to yellow in colour, and have somewhat of a “shiny” appearance compared to other aphids. One of their distinguishing features is the presence of two large, dark-green spots near the tip of their abdomen at the base of their cornicles - long tubes that look like “exhaust pipes”. These spots can easily be seen using a hand lens, especially on adults. They also have black banding on their legs and antennae, which is not present on the other common greenhouse aphid pests.

Potato Aphids

Potato Aphids are long slender aphids with antennae longer than their body. They are usually green but may be pink or red with a dark longitudinal stripe. Potato aphid’s long cornicles are light brown in color with a dark tip and are also curved outward. Potato aphids also have an indention between the base of their antennae. Aphids differ from similar insectsbecause they have two tubes called cornicles on the end of the body. Use a 10 times magnifier to see them to positively identify aphids. Root aphids resemble root mealybugs because they are covered with white wax. However, they are smaller than root mealybugs and have reduced ring like cornicles which are located on the end of their abdomen.

FEEDING DAMAGE Aphids feed by inserting their stylet-like, sucking mouthparts directly into the phloem and removing plant sap. When high aphid populations develop, plants may become stunted with curling and twisting of the young leaves. As aphids feed, a sugary plant sap, known as “honeydew”, is excreted. Honeydew promotes the growth of black sooty mold fungi which can then reduce photosynthesis. As aphids molt, their whitish cast skins may also detract from the aesthetic quality of many crops. Growers may mistake these shed skins with whiteflies. Occasionally ants may be associated with aphid-infested plants.

TRANSMISSION OF VIRUSES

In agricultural production, aphids are responsible for the transmission of a number of plant-infecting viruses. In the greenhouse, direct feeding damage is generally of more concern than virus transmission. However, aphids have been reported to transmit cucumber mosaic virus which can cause flower break and distortion on cyclamen, lisanthus and vinca.

LIFE CYCLE OF APHIDS Most types of aphids found in greenhouses do not mate. All of the aphids present are females which can give birth to live nymphs. There is no egg stage. An adult female may live for up to one month. During this time, she may give birth to 60 to 100 live nymphs. Migratory winged aphids may appear when the colony becomes overcrowded or when the food supply is depleted so they can find a new food source. Outdoors, aphids overwinter in the egg stage.

PREVENTION Inspect incoming plant material and cuttings for signs of aphids. Many aphid outbreaks occur when herbaceous perennials are introduced into the greenhouse from the overwintering cold frames. Aphids may also be carried inside on clothing or blown into the greenhouse through doors or vents. Aphid-infested weeds under the benches are frequently a source of recurring aphid problems. Inspect and remove weeds promptly. Use a weed mat barrier to prevent weed growth under the benches. The use of excessive nitrogen promotes lush growth that is favorable to aphid development.

MONITORING A regular, weekly scouting program is needed to detect aphids early before populations explode. Focus on random plant inspections of susceptible crops and cultivars to detect the wingless aphid nymphs. Look for whitish-cast skins and honeydew. Green peach aphids tend to be spread more evenly throughout the crop whereas melon aphids tend to be found in isolated hot spots. Melon aphids are also less likely to form winged adults. They usually stay on the

lower leaves and along the plant stem. Foxglove aphids inject toxic saliva as they feed leading to curled and distorted leaves, and early leaf drop. Foxglove aphids also tend to drop off of the leaves so may be hard to find. Because foxglove aphids reproduce faster at 50° to 60° F than at 77°F they are more of a problem when crops are grown cool or in the spring. Look on the leaf undersides and buds of aphid-susceptible crops. Some key bedding plants prone to aphids include: ageratum, alyssum, celosia, chrysanthemum, dahlia, gerbera daisy, herbs (many types), fuchsia, hydrangea, impatiens, pansy, pepper, portulaca, primula, salvia, snapdragon, tomato, verbena and zinnia. Some key pot plants prone to aphids include: aster, dahlia, Easter lilies, mandevilla, snapdragon and poinsettia. Some key aphid-susceptible herbaceous perennials include: arabis, aubrieta, bellis, chrysanthemum, heuchera, monarda, penstemon, phlox, salvia and viola. Yellow sticky cards will only attract winged aphids that have entered the greenhouse from outdoors, especially during the spring and early summer. Or they may indicate an aphid infestation within the greenhouse that resulted in winged aphids. However, they are not a reliable indicator of population levels in the greenhouse.

MANAGEMENT OPTIONS Biological Control In outdoor production, natural enemies, including ladybird beetles, lacewings, syrphid flies, small parasitic wasps and fungal diseases, may provide a degree of control. Outdoor environmental conditions, such as wind, rain and freezing temperatures, can also reduce aphid populations. Commercially available natural enemies may include predators, parasitoids and pathogens. In general, with the exception of the aphid predatory midge, most predators are not as effective as parasitoids in maintaining aphid populations at acceptably low levels. Repeated inundative releases of natural enemies are often needed in order to keep pace with the aphids’ high reproductive rate in the greenhouse. The goal of these repeated releases is an immediate suppression of the pest as compared to the establishment of a self reproducing population (inoculative releases). Aphids reproduce rapidly so hot spots of aphid activity may occur

and need to be treated with an alternative pesticide that is compatible with the particular natural enemies released. Consult with your supplier or check the side effects databases online. Predators Predators consume much prey during their lifetime. A predatory midge, Aphidoletes aphidimyze, can feed on more than 60 different species of aphids. This bright orange larva kills aphids by biting their knee joints, injecting a paralyzing toxin and then sucking out their body fluids. Aphidoletes is shipped as pupae in moist vermiculite; adults will emerge in 7 days at 72 °F. Adults are short-lived and tend to be active at night, so are rarely seen. After mating, a female predatory midge can lay up to 250 eggs over a 10 day period. Eggs are laid among the aphids, and hatch into larvae after 2 to 3 days. Larvae feed for 3 to 5 days. Larvae drop to the ground to pupate, so sawdust, peat or holes in the weed mat barrier on the ground are needed to provide pupation sites. This midge is most effective in the summer. It will go into diapause (a resting period of inactivity) during cool, short days (less than 8 hours of daylight). This can be prevented by the use of low intensity lights during the short days. Greenhouse temperatures should be between 60 and 80° F and 50 to 85% relative humidity. Release rates will vary with the crop grown but should be made at the first sign of aphid damage and repeated weekly until predators establish. Apply Aphidoletes in the early morning or evening near aphid colonies. Aphidoletes aphidimyza may be used preventively in combination with aphid parasitoids. Because you rarely see the adults, look for fed upon aphids (shriveled brown or black) as evidence of their activity. The green lacewing (Chrysopa rufilabiris) adults feed on nectar, pollen and honeydew. They are usually active at night. Larvae (also known as “aphid lions”) feed on aphids, thrips, spider mites and whiteflies, mealybugs and other soft bodied insects. They are commercially available as eggs or larvae. The adult can lay up to 600 eggs. Larvae will emerge from eggs in 3 to 4 days at 75 to 80° F and pupate about 14 days later. Adults emerge in 5 days. Eggs are laid individually at the end of hair like filaments on plant foliage. Larvae are quicker acting than eggs and aphid populations may be reduced in about two weeks. Ideal temperature is between 75-79° F with the minimum in temperature

for activity of 60° F. Repeat applications are often needed. The larvae tend to hide in the plant canopy during the day, but you can easily see their eggs on the foliage. A lady beetle (Hippodamia convergens) feeds on many different types of aphids and other soft bodied insects. Eggs are laid near prey and the larvae may consume from 500 to 1000 aphids. Lady bird beetles are difficult to establish in the greenhouse. Although they are inexpensive and can be easily stored in the refrigerator, they are additional concerns with their use. Hippodamia convergens are collected from the wild, so you are depleting lady bird beetles from their natural habitat. Some lady bird beetles may have been parasitized by a small wasp (Perilitus coccinellae) that as they leave the greenhouses, could potentially infect native ladybird beetles. Parasitoids Parasitoids develop in a single host and kill the host as they grow and mature. Different Aphidiusspeciesarecommercially available. Aphidius lays its eggs in aphids and the larvae develop within the aphid. The aphid is then killed as the developing larvae feed upon it. The swollen exoskeleton of the aphid remains, which is referred to as an “aphid mummy.” As the adults emerge from this mummy, you can see the small round exit hole. A. colemani is a parasitic wasp that attacks both green peach aphid and melon aphid. A related species, A. matricariae, attacks the green peach aphid. A. ervi, is used against larger species, such as the foxglove and potato aphids. Aphelinus abdominalis is a parasitic wasp that is used to control larger species of aphids such as the potato aphid. Suppliers also offer a mix of different species if you have a number of different aphid species present or are unsure of their identity. Aphid Banker Plant Systems Aphid Banker plant systems are a mini rearing system supplying a non pest species of aphid (a cereal aphid known as the bird cherry oat aphid (Rhopalosiphum padi) grown on grain plants (barley, wheat or oats) for the Aphidius colemani populations to build up and disperse thru the greenhouse. Many growers have been using screened cages for their aphid banker plants. Some have been experimenting with using hair nets placed directly over the containers of banker plants. But, some growers have mentioned that

the holes in hair nets are too large, allowing the parasitic wasps (Aphidius colemani) to contaminate the aphid population too soon, before the cereal aphid populations have build up. To solve this problem, Lloyd Traven from Peace Tree Farms, says he uses “Keystone Adjustable caps” which have a tighter weave and he doubles the net. He says they cost about .30 per banker plant. A. colemani will cluster on areas where banker plants are grown trying to get to the aphids so he uses compressed air blasts to remove them for a while to get in to work with plants. Pathogens Several types of entomopathogenic or insect-killing fungi have been developed for use against greenhouse pests. Two commercially available types are Beauveria and Paecilomyces. Beauvaria bassiana is a common soil borne fungus that occurs worldwide. Fungal spores (conidia) infect the insect through the insect’s cuticle. The fungus secretes enzymes which dissolve the insect’s cuticle. After it enters the insect’s body, the fungus produces a toxin that weakens the insect’s immune system. This fungus requires high relative humidity and moderate temperatures (65-75° F) for 8 to 10 hours in order to be effective. Beauveria may not be compatible with the convergent ladybird beetle (Hippodamia convergens) depending upon the concentration of spores applied. Thorough spray coverage is needed so that the fungal spores contact the targeted insect pest and begin the infection process. Repeated applications (three to five) may be needed for effective control. Paecilomyces fumosoroseus is a naturally occurring fungus found in infected and dead insects and some soils. The fungus infects the insect by penetrating the outer layer (cuticle) of the insect. It works best at temperatures between 72 and 86°F and high humidity. Insecticides Aphids are difficult to control with insecticides for a number of reasons. Control failures may be due to poor spray techniques, inadequate coverage or high pH in the spray tank. If aphids are present on flowers, systemic insecticides won’t be able to move into the flowers. Aphids may be difficult to reach if they are on the underside of the lowest leaves (common with

foxglove aphids). Among green peach aphid populations, resistance to organophosphates, carbamates and pyrethroid insecticides has been reported. Winged forms of the melon aphid are more resistant to organophosphate pesticides than wingless forms. Different strains of aphids may be resistant to different materials. Use pest-infested plants as indicators to monitor the effectiveness of treatments in your individual situation. Systemic materials may be more effective because aphids tend to ingest large quantities of plant sap, especially if applied before plants are in flower. Thorough coverage of the underside of leaves is needed for contact materials. Two applications of contact sprays may be more effective than one treatment.

REFERENCES Casey, C. Ed. 1999. Integrated Pest Management for Bedding Plants. A Scouting and Pest Management Guide. Cornell Cooperative Extension Pub. No. 407, 109 p. Gilrein, D. G. 2015. Time to Think About Aphids - Again. E-Gro Alert. 4(8) February 2015. Hoffman, M. and J. Sanderson. 1993. Melon Aphid. Cornell Cooperative Extension Factsheet. No. 750.50 1 p. Jandricic, S. and J. Sanderson, 2011. Early Season Pest Threat. Greenhouse Canada, 12-14. Malais, M. And W.J. Ravensburg. 2003. Knowing and Recognizing: The biology of glasshouse pests and their natural enemies. Koppert Biological Systems. 288 pp. Sanderson, J. and S. Jandricic. 2016. Out-foxing the Foxglove Aphid. GrowerTalks. October 28, 2016. Thomas, C. Greenhouse IPM with an Emphasis on Biocontrols. Publication No. AGRS-96. 89 pp. Pennsylvania Integrated Pest Management Program.

9: Cultural Practices FUNDAMENTALS OF SOIL FERTILITY Plants need 17 different minerals to grow. Carbon (C) and oxygen (O) they get from carbon dioxide in the atmosphere. Hydrogen they get from water (H2O). The other 14 elements are typically picked up from the soil. Managing soil sustainably so that it remains agriculturally productive for the long haul is a function of the biological, physical and chemical characteristics of soil, as well as dependent on adding amendments to maintain adequate levels to replace crop removal. Six of the nutrients crops get from soil are needed in relatively large amounts and are called macro nutrients. These include nitrogen (N), phosphorus (P), potassium (K), magnesium (Mg), calcium (Ca), and sulfur (S). Availability of some nutrients, for example phosphorus (P), is simply a matter of which chemical forms are present in the soil. Other nutrients, for example nitrogen (N), are very dependent on biological activity. These will be discussed in later sections. Some nutrients are commonly found in the soil as charged elements or compounds called ions.

CATION EXCHANGE CAPACITY AND BASE SATURATION Cation exchange capacity (CEC) is a measure of the soil’s ability to retain and supply nutrients, specifically the positively charged nutrients called cations. These include the base cations calcium (Ca++), magnesium (Mg++), potassium (K+), ammonium (NH4+), and many of the micronutrients. Cations are attracted to negatively charged surfaces of clay and organic particles called colloids. CEC is reported as milli-equivalents per 100 grams of soil (meq/100 g) or as centimoles of charge per kilogram (cmole+/kg). CEC can

range from below 5 meq/100 g in sandy soils low in organic matter to over 20 meq/100 g in finer textured soils and those high in organic matter. Low CEC soils are more susceptible to cation nutrient loss through leaching, and may not be able to hold enough nutrient cations for a whole season of crop production. The cations calcium (Ca++), magnesium (Mg++), potassium (K+), hydrogen (H+) and aluminum (Al+++) account for the vast majority cations adsorbed on the soil colloids in New England soils. Both H+ and Al+++ are considered acidic cations because they tend to lower soil pH while Ca++, Mg++, and K+ are considered basic cations and have little to no influence on soil pH. If all the cations are basic and none are acidic, there would be a 100% base saturation and the soil pH would be close to 7 or neutral. In acid soils there are acidic cations adsorbed on the soil colloids (called exchangeable acidity) and the percent base saturation is less than 100. A soil with a pH between 6.5 and 6.8 will typically have a base saturation of 80 to 90%.

SOIL ACIDITY, PH, AND LIMING One of the most important aspects of nutrient management is maintaining proper soil pH, which is a measure of soil acidity. A pH of 7.0 is neutral, less than 7.0 is acidic, and greater than 7.0 is alkaline. Most New England soils are naturally acidic and need to be limed periodically to keep the pH in the range of 6.5 to 6.8 desired by most vegetable crops. Scabsusceptible potato varieties are an exception but, even here, some lime may be needed to maintain the recommended pH of 5.0 to 5.2. When the soil is acidic, the availability of nitrogen (N), phosphorus (P), and potassium (K) is reduced and there are usually low amounts of calcium (Ca) and magnesium (Mg) in the soil. Under acidic conditions, most micronutrients are more soluble and are therefore more available to plants. Under very acidic conditions aluminum (Al), iron (Fe), and manganese (Mn) may be so soluble they can reach toxic levels. Soil acidity also influences soil microbes. For example, when soil pH is low (below 6.0), bacterial activity is reduced and fungal activity increases. Acidic soil conditions also reduce the effectiveness of some pesticides. The most effective way to manage soil acidity is to apply agricultural

limestone. The quantity of lime required is determined by the target pH (based on crops to be grown) and the soil’s buffering capacity. Buffering capacity refers a soil’s tendency to resist change in pH. Soil pH is only a measure of active acidity, the concentration of hydrogen ions (H+) in soil solution. It is an indicator of current soil condition. When lime is added to a soil, active acidity is neutralized by chemical reactions that remove hydrogen ions from the soil solution. However, there are also acidic cations (H+ and Al+) adsorbed on soil colloids (the CEC) which can be released into the soil solution to replace those neutralized by the lime. This is called reserve acidity. Soils such as clays or those high in organic matter have a high cation exchange capacity (CEC) and a potential for large amounts of reserve acidity. These soils are said to be well buffered. To effectively raise the soil pH, both active and reserve acidity must be neutralized. Soil test labs determine buffering capacity and lime requirement by measuring or estimating the reserve acidity. This is typically accomplished by equilibrating the soil with a buffered solution and measuring pH (buffer pH). Some laboratories calculate lime requirement from pH and soil texture (estimated CEC and base saturation) while others make this determination based on extractable aluminum levels. The neutralizing power of lime is determined by its calcium carbonate equivalence. Recommendations are based on an assumed calcium carbonate equivalence of 100. If your lime is lowerthen 100, you will need to apply more than the recommended amount, and if it is higher, you will need less. To determine the amount of lime to apply, divide the recommended amount by the percent calcium carbonate equivalence of your lime and multiply by 100. Your supplier can tell you the calcium carbonate equivalence of the lime you are purchasing. Wood ash is another amendment that may be used to manage soil acidity. The calcium carbonate equivalence of wood ash is typically around 50%, but it can vary widely. If purchasing wood ash from a supplier they will provide a recent analysis. Otherwise the wood ash should be submitted to a lab offering lime analysis to determine the calcium carbonate equivalence. The speed with which lime reacts in the soil is dependent on particle size and distribution in the soil. To determine fineness, lime particles are passed through sieves of various mesh sizes. A 10-mesh sieve has 100 openings per square inch while a 100-mesh sieve has 10,000 openings per square inch.

Lime particles that pass through a 100-mesh sieve are very fine and will dissolve and react rapidly (within a few weeks). Coarser material in the 20- to 30-mesh range will react over a longer period, such as one to two years or more. Agricultural ground limestone contains both coarse and fine particles. About half of a typical ground limestone consists of particles fine enough to react within a few months, but to be certain you should obtain a physical analysis from your supplier. Super fine or pulverized lime is sometimes used for a “quick fix” because all of the particles are fine enough to react rapidly. Lime will react most rapidly if it is thoroughly incorporated to achieve intimate contact with soil particles. This is best accomplished when lime is applied to a fairly dry soil and disked in (preferably twice). When spread on a damp soil, lime tends to cake up and doesn’t mix well. A moldboard plow has little mixing action, therefore, disking is preferred. Besides neutralizing acidity and raising soil pH, lime is also an important source of Ca and Mg for crop nutrition. It is important to select liming materials based on Ca and Mg soil content with the aim of achieving sufficient levels of each for crop nutrition. If the Mg level is low, a dolomitic lime (high magnesium lime) should be used; if Ca is below optimum a calcitic (low magnesium lime) should be used.

PLANT NUTRIENTS An element is considered essential to plant growth if it becomes part of plant tissue or is involved in metabolic functions and the plant cannot complete its lifecycle without it. There are 17 elements currently considered essential to plant growth. Listed in order of abundance in plant tissue, the 17 essential elements are: Carbon (C), Hydrogen (H), Oxygen (O), Nitrogen (N), Potassium (K), Calcium (Ca), Magnesium (Mg), Phosphorus (P), Sulfur (S), Chlorine (Cl), Iron (Fe), Boron (B), Manganese (Mn), Zinc (Zn), Copper (Cu), Molybdenum (Mo), and Nickel (Ni). Plants obtain C, H, and O from air and water during photosynthesis. Together, these three elements make up approximately 95% of a plant’s dry matter. Plants obtain the other 14 essential elements, called mineral nutrients, from soil. The mineral nutrients are classified as either macronutrients or micronutrients based on their relative abundance in plants. The six macronutrients, required in relatively large quantities, are N, P, K, S, Ca, and Mg. The eight micronutrients,

required in relatively small quantities, are Cl, Fe, B, Mn, Zn, Cu, Ni, and Mo. Other nutrients such as Silicon (Si) have shown to be beneficial in crop growth and disease suppression, but are not essential for the plant to complete its life cycle.

NUTRIENT RECOMMENDATIONS The “Plant Nutrient Recommendations” tables in each crop section can be used to determine nutrient needs based on soil test results. Nitrogen recommendations are based primarily on crop needs and are discussed later in this section under “Nitrogen Inputs.” Phosphorus and potassium recommendations are based on soil test results in relation to crop needs. In general, the goal should be to maintain nutrient elements within the optimum range as reported on the soil test. When nutrient levels are within this range, the needs of most crops will be met. If levels are low or medium (below optimum), most crops would benefit by adding the appropriate nutrient(s) to increase levels to high or optimum. However, if levels are at above optimum or very high levels, there will be no additional benefit and excess levels may reduce crop yield or quality and may cause environmental harm. This happens in fields where soil testing was not used to monitor fertility levels or when nutrients are applied even when soil levels are sufficient. When a nutrient is above optimum levels it should not be included in any amendments until the excess is taken up by crops. It may be wise to temporarily stop applying compost until nutrient levels are in the desired range. This is a practical way to manage nutrient levels if small to moderate amounts of mixed crops are to be grown. If a significant acreage of a particular crop is to be grown, fertilizers should generally be tailored to the specific needs of that crop, based on the amounts of nutrients that the crop is expected to remove during the growing season. If the soil tests indicate that a nutrient is high/optimum it is likely that the soil will supply enough to meet the crop’s needs. However, many growers will apply enough of the nutrient to replace what is removed by the crop. If the test level is above optimum/ very high, additional applications should normally be avoided unless the crop has an unusually high demand for a specific nutrient. Occasionally, nutrient applications may exceed the soil test recommendation or the expected average removed by the crop because a

particular cultivar is considered a heavy feeder such as long season Russet potatoes. Or, for example, a large crop of tomatoes can be expected to remove a large amount of potassium and it may be justified to apply some of this nutrient even if the soil test indicates a level somewhat above high/ optimum. The nutrient recommendation tables for each crop have been developed on this basis. This can also be a practical way to determine nutrient needs of high value crops, even when they are grown on a small scale. It is important to keep in mind that factors other than nutrients may limit crop potential, and simply adding more nutrients will not solve such problems. Nitrogen Nitrogen is essential to nearly every aspect of plant growth, but it is one of the most difficult nutrients to manage. When plant available N exceeds crop demand, nitrate (NO3) accumulates in soil increasing the risk of N loss to the environment. Excessive levels of available N can also produce succulent plants that are more susceptible to environmental stress and pest pressure. When plant-available N is too low, crop health and productivity suffer. The key to successfully managing N is to determine the relatively narrow range between too much and too little - this is not an easy task. Having an understanding of the forms of N in the soil and the factors that influence its behavior will help improve management of this dynamic nutrient. The Nitrogen Cycle Practical knowledge of the N cycle is key to effective and efficient N management. The N cycle is extremely dynamic and, as illustrated in Fig. 9.1, its behavior in soil is complex. Nitrogen transformations and losses are affected by the form of N added, soil characteristics and conditions, and the vagaries of the weather. The rate and magnitude of N transformations and losses are difficult to accurately predict. Nitrogen Inputs As can be seen from the N cycle, there are two forms of the N used by plants: ammonium (NH4) and nitrate (NO3). In addition to commercial fertilizer sources, available N may be added to the soil through mineralization (the microbial conversion of organic N to ammonium and then nitrate) of soil

organic matter, manure and other organic residuals.

Fig. 9.1. The Nitrogen Cycle

Nitrogen in soil organic matter Organic matter contains the largest pool of soil N, usually comprising more than 90 percent of total soil N. The total amount of organic matter N in the plow layer of agricultural soils is impressively large. As a rule of thumb, you can assume that for each 1% of organic matter in the surface 6” or 7” of soil, there are 1000 lbs of N per acre. Thus, a soil with 3% organic matter contains about 3000 lb of N per acre. The amount of total organic matter N that becomes available to plants in any one year, is relatively small as a percentage of the total organic matter, but can be large in some years relative to the amount of N needed for plant growth. Research has shown that for most soils 2% to 4% of the total organic matter N is converted (mineralized) annually to forms plants can use. This is roughly equivalent to 20 to 40 lb of available N per acre for each 1% of organic matter in the surface 6” or 7” of soil. A soil with 8% organic matter content, therefore, will mineralize, or make available to plants, 160 to 320 lb N/acre from the organic matter. This amount of N would be sufficient for most vegetable crops in a relatively dry year with little leaching or denitrification. This mineralization is not constant throughout the growing season. A flush of available N is mineralized in late spring with lower rates of mineralization occurring during the season. Moisture conditions will greatly influence the mineralization during the season, with high rates when the soil

is near water holding capacity in a well aerated soil, and lower rates when the soil is dry. Small flushes of N will be mineralized when soils are re-wetted during the season. The rate of mineralization is dependent on microbial activity, especially bacterial activity. Such activity is favored by warm, well aerated soils with adequate, but not excessive moisture and a pH above 6.0. These conditions are also favorable for the growth of most vegetables. Nitrogen in manures and other waste products The N content of manures is highly variable. Differences in N content are due to the species of animal, the animal’s age and diet, the moisture content of the manure, handling and storage, and the amount of bedding in the manure. The N fertilizer equivalent of a manure varies not only with the total N content of the manure, but also with the timing and method of manure application. The values in Table 1 are based on numerous analyses of Connecticut manures as well as published data from other states. If specific manure analysis data for the farm are not available, growers should estimate N credits using these or other book values. The time elapsed between spreading and incorporation of manure is also important. About half of the N in dairy manure and three quarters of the N in poultry manure is in the form of ammonia (NH3) which is volatile. The following manure application methods are listed in order of being most effective to least effective for reducing ammonia volatility: Tine injection, disc injection, immediate incorporation after surface application, band spreading with trailing hoses or trailing shoe without incorporation, broadcast application with incorporation. Broadcast application of slurry manure without incorporation should be avoided at all times because this method increases air to ammonia contact and allows time for all ammonia to be lost. Research has shown that in reduced or no-till fields where manure must be surface applied without incorporation, ammonia can be conserved if applied during cold temperatures, low wind speeds and especially to a growing cover. A growing cover also reduces manure run-off or leaching. Previous manure applications Up to 50% of the total N in cow manure is available to crops in the year of application. Between 5% and 10% of the total N applied is released the year after the manure is added. Smaller amounts are furnished in subsequent years. The quantity of N released the year after a single application of 20 tons

per acre of cow manure is small (about 15 lb N per acre). However, in cases where manure has been applied at high rates (30 to 40 tons per acre) for several years, the N furnished from previous manure increases substantially. The buildup of a soil’s N-supplying capacity resulting from previous applications of manure has important consequences for efficient N management, two of which are: 1) The amount of fertilizer N needed for the crop decreases annually; and 2) If all the crop’s N needs are being supplied by manure, the amount of manure needed decreases yearly. Table 9.1: Nitrogen Credits from Manure Incorporated Before Planting

1 Dry matter.

With caged layer poultry manure, a higher percentage of the total N in the manure is converted to plant-available forms in the year of application. Consequently, there is relatively less carry-over of N to crops in succeeding years. This is due to the nature of the organic N compounds in poultry manure. This does not mean, however, that there is never any carry-over of N from poultry manure applications. If excessive rates of poultry manure (or commercial N fertilizers) are used, high levels of residual inorganic N, including nitrate (NO3), may accumulate in soil. High levels of soil nitrate in the fall, winter and spring have the potential to pollute groundwater and coastal seawater. Previous crops: Many vegetables leave little residue in the field and thus they provide little N benefit to subsequent crops. However, previous forage or cover crops can supply appreciable amounts of N to succeeding

crops. Legumes, such as alfalfa and red clover, can furnish 100 lb or more of N to crops that follow (Table 9.2). Other legumes, mixed grass-legume stands and grass sods supply less N to succeeding crops. Keep in mind that most of the N is in the leaves, not the roots. If a legume hay crop is harvested, most of the N is removed from the field along with the hay.

Table 9.2: Nitrogen Credits for Previous Crops Previous Crop

Nitrogen CreditLb N per acre

“Fair” clover (20-60% stand)

40 to 60

“Good” clover (60-100% stand)

60 to 90

“Fair” alfalfa (20-60% stand)

60 to 90

“Good” alfalfa (60-100% stand)

100 to 150

“Good” hairy vetch winter cover crop

120 to 150

Grass sod

20 to 40

Sweet corn stalks

30

Synthetic fertilizers Fertilizers used to supply N include urea (46-0-0), diammonium phosphate (DAP: 18-46-0), monoammonium phosphate (MAP: 11-48-0), ammonium nitrate (not readily available), urea-ammonium nitrate solution (UAN: 32-0-0), calcium ammonium nitrate (this has generally replaced ammonium nitrate), calcium nitrate, potassium nitrate and various manufactured and blended fertilizers such as 15-8-12, 15-15-15 and 10-1010. In bulk blended or custom blended mixes, N-containing fertilizers with almost any grade can be provided. Nitrogen Losses Nitrogen losses occur in several ways. The loss of available soil N not only costs growers money, it has the potential to negatively impact both air and water quality. Understanding the cause of N losses can help growers make management decisions to improve N use efficiency and minimize negative environmental impact. Volatilization Losses These losses occur mainly from surface applied manures and urea. The losses can be substantial; more than 30% of the N in topdressed urea can be volatilized if there is no rain or incorporation within two or three days of

application. Losses are greatest on warm, breezy days. Volatilization losses tend to be greater from sandy soils and when pH values are above 7.0. Delaying the incorporation of manures after they are spread also leads to volatilization losses of N. Under the right conditions more than 50% of the ammonium N may be volatilized within the first 48 hours following surface application of manure without incorporation. Not only does volatilization reduce the fertilizer value of manure and urea, it can degrade air and water quality. Ammonia in the atmosphere can form particulates that contribute to smog. Ammonia emissions can also contribute to eutrophication of surface waters via atmospheric deposition. Leaching Losses The nitrate form of N is especially mobile in soil and is prone to leaching losses. Leaching losses are greatest on permeable, well-drained to excessively-drained soils underlain by sands or sands and gravel when water percolates through the soil. Percolation rates are generally highest when the soil surface is not frozen and evapotranspiration rates are low. Thus, October, November, early December, late March and April are times that percolation rates are highest and leaching potential is greatest. This is why nitrate remaining in the soil after the harvest of annual crops such as corn in September is particularly susceptible to leaching. The use of cover crops following cash crops can take up this residual N and prevent it from leaching. The N will then be released for crop use after the cover crop is plowed down in the spring. Of course, leaching can occur any time there is sufficient rainfall or irrigation to saturate the soil. This is why it is important to attempt to match fertilizer N application rates with crop N needs. Nitrate leaching accounts for the vast majority of N losses from cropland. Nitrate leaching can have a direct impact on water quality. When nitrate leaching contaminates groundwater serving drinking water supplies, human health can be impacted. The greatest concern is for infants; high levels of nitrate can be toxic to newborns, causing anoxia also known as “bluebaby” syndrome. High nitrate levels in drinking water are also harmful to young or pregnant livestock. Depending on regional hydrology, leaching losses of nitrate can also contaminate surface waters causing eutrophic conditions. Denitrification Losses

These losses occur when nitrate is converted to gases such as nitrous oxide (N2O) and nitrogen (N2). The conversions occur when the soil becomes saturated with water. Poorly drained soils are particularly susceptible to such losses. In especially wet years on some soils, more than half the fertilizer N applied can be lost through denitrification. The most favorable conditions for denitrification tend to occur in early spring and late fall. Minimizing the concentration of nitrate in the soil during these periods by delaying N application in the spring and planting cover crops in the fall will help reduce denitrification losses. Most of the N lost during denitrification is in the form of the inert nitrogen gas (N2) which has no negative impact on the environment (our atmosphere is approximately 78% N2). Only a small percentage of denitrified N is lost as nitrous oxide (N2O); however, it is a powerful greenhouse gas. The impact of 1 pound of nitrous oxide on atmospheric warming is over 300 times greater than 1 pound of carbon dioxide. Agricultural activities account for over 70% of nitrous oxide emissions in the US. Immobilization Immobilization occurs when soil micro-organisms absorb plant-available forms of N. The N is not really lost from the soil because it is held in the bodies of the microorganisms. Eventually, this N will be converted back to plant-available forms. In the meantime, however, plants are deprived of this N, and N shortages in the plants may develop. Immobilization takes place when highly carbonaceous materials such as straw, sawdust or woodchips are incorporated into the soil. Manure with large amounts of bedding and compost with C:N ratios greater than 30:1 may cause some immobilization. Crop Removal of Nitrogen A significant quantity of N is removed soil via crop harvest. For example, good sweet corn crops may remove over 150 lb of N per acre annually. Anticipated crop removal of N is one of the factors used in making N fertilizer recommendations. Depending on the crop, variable amounts of the N taken up by the crop are returned to the soil after harvest in nonharvested plant parts. With sweet corn this can be as much as 100 lb N/A. As these leaves and stalks decompose, the N is released into the soil for use by a subsequent crop. Cover crops can take up much of this N preventing it

from being lost via leaching or denitrification.

NITROGEN MANAGEMENT Topdressing and Sidedressing Nitrogen Topdressing is defined as a fertilizer application to a crop any time after planting. In popular usage, topdressing sometimes refers to a broadcast application of fertilizer made after planting. However, the fertilizer can be sidedressed as a band along the side of the row of a growing crop. Sidedressing is commonly done immediately before or during cultivation. When urea containing fertilizers are used, cultivation helps reduce volatilization losses. Sidedressing of relatively soluble N fertilizer is an important component of efficient nitrogen management. The N accumulation pattern for annual crops is very similar to biomass accumulation. Early in the season, when crop growth is slow, crop N needs are very small. A starter fertilizer is generally sufficient to satisfy those needs. Any soil nitrate in excess of crop N needs during this period is prone to leaching and/or denitrification losses. The next phase of crop development is characterized by rapid vegetative growth. The N demand during this phase is the highest of the growing season. As much as 85% of the total N uptake occurs during this period. Efficient recovery of fertilizer N can be achieved by sidedressing fertilizer N immediately before this phase. Delaying application of a large portion N fertilizer until sidedress also allows growers to use the Pre-sidedress Soil Nitrate Test (PSNT) to help determine N needs. Pre-sidedress Soil Nitrate Test (PSNT) The dynamic nature of the N cycle and its sensitivity to weather limits the value of routine, pre-season soil testing for predicting N availability during the season in our humid environment. However, under certain circumstances, in-season soil testing has proven useful. The PSNT, developed by Dr. Fred Magdoff at the University of Vermont in the early 1980s, was originally intended to help estimate the amount of available N for field corn in fields where manure had been applied and/or forage legumes were grown in rotation. Over the last thirty years research conducted in the Northeast has shown the PSNT useful for improving N management of

several vegetable crops including sweet corn, peppers, pumpkin, winter squash, and cabbage. The PSNT is most suitable for use with annual crops, which accumulate N rapidly within a single growing season. The PSNT is especially useful where large amounts of N from mineralization are expected, and the test works best when pre-plant and starter fertilizer N rates are less than about 50 lbs N per acre. PSNT samples are collected about a week before the rapid growth phase, to provide an indication of how much N has been made available from mineralization. During wet springs with heavy leaching rains, or in sandy soils with rapid losses, the PSNT will also provide some indication of how much N remains in the root zone. As with all soil testing, information from a PSNT should be used along with the grower’s experience and knowledge of the field. Interpretation of the PSNT is also crop specific. Research in the Northeast has shown that when the soil nitrate N level is above 20 to 25 ppm there is rarely an economic response to the application of sidedress fertilizer N for sweet corn. Based on research and experience, a threshold of 25 to 30 ppm seems appropriate for peppers, tomatoes, butternut squash, cabbage, pumpkin, and probably other long-season vegetable crops. When PSNT values are below threshold levels, the appropriate rate of sidedress N should be determined based on the level of nitrate N reported, previous N application, realistic yield expectation, the field’s management history, and growing season conditions. See Table 9.3 for recommendations on timing of sampling and making sidedressing applications of N based on PSNT for many vegetable crops. Samples for the PSNT should consist of a well-mixed composite of 10 to 20 cores or slices of soil to a depth of 12”. This is a deeper sample than what it is recommended for routine soil sampling. A deeper sample is required for nitrate testing to accurately reflect the concentration in the effective root zone due to its mobility in soil. Avoid sampling fertilizer bands or areas that may have received extra N. About one cup of the composite should be dried to stabilize the nitrate. A good method is to spread a thin layer of the soil on a cookie sheet or aluminum foil to air dry. Use a fan to reduce drying time. Do not place damp samples on absorbent material because it can absorb some of the nitrate. You can skip the drying step if you can deliver the samples to the soil testing lab in less than 24 hours; however, samples should be kept cool. Fields should be sampled for the PSNT about a week before the time when

sidedressing is normally done. This should allow adequate time for drying, shipping, and testing (turnaround time in the lab is about 24 hours) and for you to plan your fertilizer program.

Table 9.3. Timing of PSNT and sidedress nitrogen needs of crops

a If soils have 0-30 ppm nitrate, apply the full sidedress amount recommended. For sweet corn, the threshold is 25 ppm nitrate. Above 30 ppm no additional N is needed and could hurt yields.

Phosphorus Phosphorus (P) is referred to as phosphate, or P2O5, for the purposes of fertilizer grades and recommendations. We don’t apply P in this form, but it has become the standard unit of measure over many years. The amount of extractable P in a soil should not exceed the optimum soil test range to obtain the most economic return to P applications. Extractable P should not exceed the Environmental critical concentration, which is much higher than the Optimum range, to maintain water quality. When extractable P exceeds the Environmental critical concentration, the risk of dissolved P loss in subsurface water flow or runoff in amounts that pollute surface water

(especially lakes and ponds) is significantly increased. The pollution occurs from the P stimulating excessive growth of algae in lakes and ponds. When the P levels are reduced by uptake by the algae, the algae die and their biomass is rapidly decomposed by micro-organisms, which can reduce oxygen levels below the level needed by fish and shellfish, resulting in large die offs of aquatic life. Excessive P amounts in soils are difficult to reduce because vegetable crops remove little P from the soil compared to N or K. For example, sweet corn takes up about 155 lb/A of N and about 105 lb/A of K, but only about 20 lb/A of P. However, growers commonly apply about 100 lb/A of P annually. This is justified only if soil test P levels are below optimum, because amounts of P greater than crop uptake are required to increase soil test levels from below optimum into the optimum range. However, if the soil test level for P is above optimum, there is little if any crop response to additional P applications. Plant uptake of P is extremely slow in cold soils. For this reason, when planting early into soils testing Optimum or lower, it is often advisable to apply up to 30 pounds of P2O5 as starter fertilizer in a band about 2” below and 2” to the side of the seed when planting, or as a liquid around transplants. Keep in mind that P availability is reduced in alkaline soils (pH >7.3) as it will bind with Ca becoming unavailable to plants and in acidic soils (pH 50%) nitrate fertilizers. However, these fertilizers also have elevated trace element levels which may raise iron (Fe) and manganese (Mn) to toxic levels at low pH. Both are acid-forming fertilizers, but 20-10-20 has the greater potential acidity. 15-15-15 Geranium Special. “Triple 15” is a good alternative to the Peat-Lite Specials for crops sensitive to trace element toxicities. Trace element levels supplied by this fertilizer are lower than the Peat-Lite Specials. Otherwise, at the same rate of nitrogen (N), plant response will be very similar to 15-16-17. This is an acid-forming fertilizer also; the potential acidity is slightly greater than 15-16-17. 20-20-20 General Purpose. Growers who use this fertilizer on soilless media risk ammonium toxicity because the nitrogen in this fertilizer is 75% ammonium and urea. Some growers who use media containing soil do not appear to have problems. If 20-20-20 is used, the growing medium should be tested frequently for ammonium. 20-20-20 supplies trace elements and has the greatest potential acidity of fertilizers commonly used in New England greenhouses. Tomato, eggplant and pepper plants are especially sensitive to ammonium. High ammonium levels, especially in soilless mixes, can reduce plant growth and cause yellowing of the foliage. Low Phosphorus (P) Fertilizers (20-0-20, 20-1-20, 15-0-15). These

fertilizers can be tried as an alternative to chemical growth regulators for vegetable bedding plants. This technique of growth control is sometimes called “phosphorus starvation.” It is generally believed that more P than necessary is being applied to greenhouse crops. Too much P may cause plants to stretch and P is a ground water pollutant. Unfortunately, in terms of height control, these fertilizers may be of no benefit if they are applied to a growth medium containing superphosphate or a high starter charge of P. Also, there is a risk of P deficiency if the fertilizers are used continuously with low P growth media. The low P fertilizers are quite different in many ways. 15-0-15 and 20-0-20 supply Calcium (Ca). 15-0-15 is a basic (raises pH) fertilizer containing about 95% nitrate and 20-0-20 is a neutral fertilizer and is 50% nitrate. 20-1-20 is an acidic fertilizer and it does not supply Ca, but it is about 70% nitrate. Calcium nitrate and potassium nitrate (15-0-15). Use of this fertilizer combination greatly reduces the chance of trace element toxicities. Some growers alternate its use with the Peat-Lite Specials on a 2-3 week basis to supply Ca and to counter the acidic effect of the Peat-Lites. However, both superphosphate and a trace element fertilizer must be incorporated in the growing medium if this combination is to be used as the sole fertilizer.

NITROGEN, PHOSPHORUS, POTASSIUM Nitrogen. Nitrogen concentration in the greenhouse fertilizer program has a greater affect on the growth of transplants in the greenhouse than either phosphorus or potassium. Increasing the level of nitrogen results in taller transplants with thicker stem diameters and heavier plant weights. Applying too much nitrogen in the greenhouse results in soft, poor quality transplants. These lush transplants may also be more prone to phloem feeding insects such as aphids and to foliar blights. Phosphorus: Phosphorus has a limited affect on the growth of bedding plants when compared to nitrogen, but should be included as part of a complete fertilizer. Increasing the phosphorus concentration results in a moderate increase in transplant height, stem diameter, and shoot fresh and dry weight. If phosphorus is restricted to the point at which the plants show extreme phosphorus deficiency (purple leaves and stems, stunted plants), field performance will be reduced.

Potassium: Potassium has the least affect on the growth of plug tomato transplants of the three major nutrients. Adequate potassium is applied as part of a complete fertilizer.

GUIDELINES FOR RATES AND FREQUENCY OF FERTILIZER Small, slow-growing plants should receive lower rates or less frequent application until they are well-established. Care should be taken not to overfertilize vegetable bedding plants to avoid overgrown plants. Young seedlings are especially vulnerable to injury from high soluble salts. While plants are in the plug or seedling stage, use a complete water soluble fertilizer at the rate of 50 – 100 ppm N every time plants are watered and use clear water (no fertilizer) every third watering. Use the lower rate (50 ppm) early and the higher rate (100 ppm) later if the seedlings are to be held in the flat or tray three or more weeks before transplanting. Shortly after transplanting, as plants approach rapid growth, increase the rate to 200 ppm N at every watering or 300 ppm N once every 7 days, watering with clear water 2 or 3 times in-between each fertilization. Fertilizer Solution Volume: The volume of fertilizer solution applied has a dramatic affect on the growth of the vegetable bedding plants. As the volume of water-soluble fertilizer increases, the quantity of nutrients delivered to the plant also increases resulting in an increase in height, stem diameter and plant weight. Doubling the volume applied also doubles the amount of each nutrient potentially available to the plant. Plant Growth Rate and Environmental Conditions. In general, nutrient requirements of vegetable bedding plants are greatest during periods of rapid growth. Too much fertilizer during slow growth periods may lead to excess soluble salts; failure to provide enough fertilizer during periods of rapid growth will lead to nutrient deficiency.

NUTRITIONAL PROBLEMS Vegetable bedding plants and transplants are subject to the same nutrient disorders as other plants. Early in production serious problems are: high

soluble salts, trace element toxicities, and ammonium toxicity. Late in production, particularly in cell packs, plants may develop nitrogen deficiency symptoms as the earliest indication of insufficient fertility. Soluble Salts. Injury to vegetable bedding plants from excess salts seems to be most common shortly after transplanting. Seedlings are much less tolerant than established, rapidly growing plants. Some soilless mixes may contain enough “starter charge” to cause excess salts problems in the first few weeks after transplanting, particularly when a water-soluble fertilizer is also applied. Excessive drying, poor drainage, and uneven watering are factors, which can aggravate this problem. Check roots of plants often and conduct regular soil tests to identify and prevent problems. It is difficult to diagnose a soluble salts problem by symptoms alone. Often nutrient deficiencies and root diseases cause the same symptoms. Therefore, a greenhouse (not field) soil media test is advisable. Trace element toxicities. Iron (Fe) and/or manganese (Mn) can be accumulated to toxic levels by tomato plants. Symptoms appear as numerous small dark spots and mottling of the foliage. The potential sources of excess Fe and Mn are: trace element fertilizers in the mix, water-soluble fertilizers with elevated trace elements levels, and sometimes irrigation water. Low growth medium pH aggravates the problem by increasing Fe and Mn availability. Toxicity can be avoided by keeping the pH in the range of 5.8 – 6.0 for susceptible crops and by the use of fertilizers with lower trace element levels. Ammonium toxicity. This is less common today because most growers use water-soluble fertilizers that supply about 50/50 ammonium and nitrate to fertilize plants in soilless media. Tomato, eggplant, and pepper are most sensitive to ammonium nitrogen, but many other vegetable bedding plants can be harmed if ammonium becomes excessive. Too much ammonium during the early spring (February or March) in low light and cool media conditions can be toxic to plants. Plant growth may be reduced with yellowing of the foliage.

ORGANIC NUTRIENT MANAGEMENT The quality of the planting mix is important to insure proper plant health in organic production. Conventional growing media that contains synthetic

ingredients (wetting agent, chemical fertilizer) cannot be used in organic production of field transplants, and vegetable bedding plants. However, acceptable growing media can be composed from a wide variety of approved materials. These organic blends may be purchased off-the-shelf, customblended by manufacturers, or produced on-the-farm. Most commercial potting mixes contain synthetic fertilizers and wetting agents that do not meet organic standards. One alternative is to arrange a special order from a commercial supplier who agrees to exclude starter fertilizers and wetting agents and then, plan to add your own. Purchasing a commercially prepared organic mix is the easiest way to get started and most growers choose this option to reduce the risk of soil-borne diseases. Common components such as peat moss, perlite, vermiculite, and coconut coir are generally acceptable for organic certification, but check with your organic certifier. Compost, being the most renewable, is a preferred material for many organic growers. Researcher chooses a 50/50 mixture of peat and compost with a pH of 6.0 as an organic potting media. He makes his own compost from garden waste, straw, hay and sheep/horse manure and screens the compost to provide a uniform product. This is just one of many options; however the finished compost product needs to have good physical, chemical and biological properties. Since compost can vary from batch to batch, there can be variation in the performance of mixes, even when using the same recipe. If making or buying compost for your potting mix, try to have it ready at least 6 months before you need it. This allows time for the compost to mature so that nutrients are stabilized, phytotoxic compounds have degraded and disease suppressive beneficial microbes have a chance to increase. It is very important to test the compost before you use it to determine pH, available nutrients, and soluble salt levels. Organic potting mixes may contain from 20 to 50% compost by volume depending upon the crop, container size and growing conditions. 100% compost is generally not advisable. Another option recommended by Eliot Coleman, from Four Season Farm, Maine is to use a blend of peat and compost or peat-based soilless medium and compost mixed with 14 lbs. per cubic yard of equal parts blood meal or alfalfa meal for nitrogen, rock phosphate for phosphorus and green

sand or organic approved potassium sulfate for potassium. He lets the blend sit for a month or more before use. A third option is to use mature, well-balanced compost blended with peat and possibly perlite and/or vermiculite for aeration to supply all the nutrients to grow and finish transplants. Nutrient sources such as alfalfa, alfalfa meal and other organic approved components can be incorporated during the composting process. Compost will mature during the fall and can be stored for use until spring. These last two options add some nutrients to the root medium, so plant nutrition is not dependent on liquid fertilizer. However, these options require purchasing the organic nutrient sources and making sure the rate and method of application are correct. There are also many fertility management options such as supplementing with liquid organic fertilizers. Many growers are familiar with fish fertilizers made from waste products of the ocean fish processing industry. The material is a thick, heavy liquid supplying plant nutrients at presumably varying levels of availability. Fish fertilizers probably supply mostly ammonium nitrogen which could be a disadvantage for some plants. Also, fish fertilizers can be a problem to store diluted because they spoil and they can be difficult to inject through some systems. In our area the Neptune’s Harvest brand is the most commonly available fish fertilizer and it is OMRI-approved for organic greenhouses. In his studies, researcher, used liquid fish emulsion (5-1-1) as a sole fertilizer with a soilless peat based medium and coconut coir successfully for several months. Fish emulsion fertilizers are likely to have an odor and fertilizing less often may be preferred, for example, every two weeks or once a month. The rate applied will vary depending upon how often one fertilizes. Plants fertilized with organic fertilizers will not show the rapid growth response seen with synthetic fertilizers. With this method, Dr. Biernbaum suggests beginning with a growing media containing 60-70% peat and 3040% perlite and/or vermiculite without fertilizer and wetting agent, but limed to a pH of 5.5 to 6.5. Apply the water soluble organic fertilizer as needed, usually soon after seed emergence or at transplanting. Many growers are using Daniels 10-4-3 a liquid, “organically-based” fertilizer. The organic portion is oilseed extract. Most of the nutrients, however, are derived from inorganic salts and for this reason it cannot be

certified as being organic. Daniels Pinnacle 3-1-1 is a less well-known liquid fertilizer. It is an organic fertilizer, as most nutrients are derived from oilseed extract and extra nitrogen is provided by sodium nitrate (“Chilean nitrate”). This fertilizer is a great step forward in finding organic fertilizers that can be easily applied by growers using the systems to which they are accustomed. However, Pinnacle spoils after dilution and it does not provide adequate nutrition at the label rates with the result that growth is reduced and nutrient deficiency symptoms develop, likely related to high pH and iron deficiency. Pinnacle seems to work better when it is alternated with fish emulsion. There are no deficiency symptoms, pH is lower and plant nutrient status is better.

PLANTING Handling Growing Media How soilless growing media is handled can greatly influence the air space and available water for the plant roots. The major concern is to preserve the air space or porosity to insure healthy root growth. You want to prevent compaction that encourages damping off diseases and poor root growth. Containers, including plug trays, should be lightly filled and the excess media brushed off the top. At no time should any growing containers be stacked. Stacking containers causes compacted media. This damage cannot be remedied after the compaction has developed. Add water to peatbased mixes before filling plug trays to help create more aeration. If mixing your own media, thoroughly mix components, but do not over-mix.

PRODUCTION SCHEDULES Starting seeds too soon, will result in overgrown plants of poor quality. The following are guidelines for growing vegetable bedding plants. Note the number of weeks from seed to sale or transplant. This will vary according to different growing conditions and should serve only as a guide. Crop

Germination Temperature (°F)

Optimum Day Production Temperature (°F)

Minimum Night Temperature (°F)

Weeks from Seed to Sale

Weeks from Seed to Transplant in Field

Basil

70 +

70 to 75

60 to 65

4 to 7

4 to 6

Broccoli

70 to 75

65 to 70

55 to 60

4 to 7

4 to 6

Cabbage

70 to 75

65 to 70

55 to 60

4 to 7

4 to 6

Cauliflower

70 to 75

65 to 70

55 to 60

4 to 7

4 to 6

Celery

70 to 75

60

60

5 to 7

8 to 10

Chard

70

60

50

Cucumber

70 to 75

70 to 75

60 to 65

2 to 3

3

Eggplant

70 to 80

70 to 80

60

7 to 9

6 to 8

Lettuce

65 to 70

60 to 65

50

3 to 5

Leeks

75

65 to 70

55 to 60

10 12

Melons

70 to 85

70 to 75

60 to 65

2 to 3

2 to 3

Onions

75

65 to 70

55 to 60

10 12

10 to 12

Peppers

75 to 85

70 to 75

60

6 to 8

6 to 8

Squash/ Pumpkin

70 to 85

70 to 75

65

2 to 3

2 to 3

Tomatoes

75

65 to 75

60

5 to 8

5 to 8

Watermelon

80 to 90

70 to 80

65 to 70

3 to 4

3 to 4

to

to

10 to 12

Note: The greater the difference between daytime and nighttime temperatures, the more plants will “stretch” (stems elongate).

GERMINATION TIPS Warm temperatures and uniform moisture are needed to ensure successful germination and get the plants off to an even start. Many germination chamber systems are commercially available including custom built germination units. Many growers use bottom heat or root zone heating to provide warm, even temperatures. Rubber tubing or mats with hot water are placed on the bench top under the plants. A weed mat barrier is placed on the top of the bench to help spread the heat with skirts on the side to help contain the heat. In all systems, it is important to remove flats from the germination chamber as soon as radicles break through the seed coat to avoid seedling stretching. Experience and experimentation with your total seeding system is the key to uniformity and success. Celery

Celery seeds germinate best at 70 – 75°F. To prevent bolting, maintain greenhouse temperatures above 55°F Cole Crops (Cabbage, Broccoli, Brussels sprouts, Cauliflower) To prevent premature seeding or bolting, avoid exposing transplants to temperatures below 50°F for long periods (week or more). The cold temperatures cause the development of premature heads or “buttoning” in cauliflower and broccoli. Any stress or check in growth results in a “wirestem” and plants will not become well established in the field or garden resulting in reduced yields and performance. Eggplant Eggplant seed can be directly sowed into 50 cell trays to shorten the time needed to produce transplants by approximately one week. Germinate seed in flats at 70 to 75°F. Eggplants are susceptible to chilling injury and should not be grown below 40°F. Any stress or check in growth will result in tough woody stems and transplants that will have a tough time getting started later in the field or garden. Tomatoes Tomato seeds germinate best at 75°F. As soon as there is any evidence of germination, they should be removed from the mist and bottom heat. The ideal root-zone temperature is 77-86°F during the first four weeks of growth and 68 to 77°F during the fifth and sixth weeks. Optimal growing-on day temperatures are 65 to 75°F with minimum night temperatures of 60°F. Exposure of tomato plants to temperature below 60°F will likely result in rough fruit (catfacing) on the first few clusters. Transplant young seedling into 2” to 3” containers when they have two true leaves and grow on until planted in the field. For earliest production, some growers finish their transplants into 6” or larger containers. Peppers Seeds germinate at 85 to 90°F. Note that germination is very slow at lower temperatures. Seedlings develop well at 75°F daytime and 65° F night temperatures. Seeds may be directly sown into 72-cell trays for early production. Peppers are prone to damping off diseases especially if the media is compacted. They are also susceptible to transplant shock.

Vine Crops Cucurbits do not transplant well, and are best to sown in the final container. After germination, excess plants can be thinned. Transplants Transplants can be grown in all types and sizes of containers. Before sowing, one needs to decide whether germination and finishing will occur in the same container or whether seeds will be sown in one container followed by transplanting to a finishing container. Germinating and growing in small plugs requires more attention to detail and is probably best done by local, specialty propagators. Direct sowing of transplants to be finished in 200 - 128 cell or larger trays may be more manageable for many growers. Before making a decision, consider your available labor, and amount of greenhouse space, and the cost and benefit of each production method. Plug seedlings should be transplanted as soon as possible after they have reached finished size. Purchased plugs: Purchase transplants from a reputable local supplier to minimize the potential of importing severe disease and insect problems that are common in other regions of the country. Open and unpack the boxes immediately upon arrival and check the physical condition of the plants. Inspect plants for root and foliar diseases and for insects and mites. Report any damage or discrepancies immediately to your supplier (most companies want to hear within 24 hours). Photographs are also helpful. Place plant trays on benches and water thoroughly with plain water (no fertilizer); be sure that plugs on the edges of the trays are thoroughly watered. Plugs can dry out quickly due to the small volume of growing medium; check the trays 2 or 3 times daily for watering. After the initial watering, apply a general-purpose fertilizer (such as 20-10-20) at 50 to 60 ppm of nitrogen at every other watering. Allow plants to acclimate to the greenhouse conditions for 24 to 48 hours before transplanting. Transplanting to a finishing container: Water the plug trays thoroughly 2 to 3 hours before transplanting; this aids in removing the plugs from the trays. Prepare your cell packs or pots by filling them with pre-moistened growing medium and pre-dibbled holes for the plugs. Lightly fill containers

and brush off excess. To prevent compaction, do not pack down or stack (“nest”) filled flats. Take special care during transplanting to handle plants gently and avoid planting too deeply. Stems of tender seedlings can be easily injured when workers grasp or “pinch” the stems too tightly. This often leads to stem cankers causing plants to wilt and die. Plant plugs at the same depth as the original plug. Some transplants may have elongated stems and it is tempting to “bury” the stem. Resist the temptation, except for more adaptable tomato plants. See information under the specific crops for additional information on transplant production and planting.

PLANT CULTURE Proper Watering Practices Hand watering and overhead irrigation systems are the primary methods of watering vegetable bedding plants and transplants. The amount of water and frequency of watering will vary depending on container size, growing media, greenhouse ventilation and weather conditions. It is important to water thoroughly, to moisten the entire container, which will promote root growth to the bottom of the container. If this is not done, root growth will develop in the upper part of the container and plants will be more prone to drying and drought stress. Allow plants to dry down before watering, but do not let the plant wilt severely, as this will damage roots. Vegetable bedding plants should be watered thoroughly early enough in the day to allow foliage to dry before evening. If foliage remains wet overnight, foliar disease problems will develop. Limit water leaching to 10 to 20% to limit nutrient runoff. Most commercial mixes contain a wetting agent which provides initial hydration and improves wettability of the mix. Older mixes (stored longer than 8 months) are harder to wet and the addition of a liquid wetting agent may be needed. Managing Plant Height A review of pesticide labels indicates that Sumagic (uniconazole) is the only growth regulator labeled for use on a limited group of vegetable transplants (tomato, pepper, ornamental pepper, eggplant, tomatillo, ground

cherry and pepino). Apply Sumagic only as a foliar spray at a rate of 2-10 ppm. As with any plant growth regulator, it is recommended to test growth regulator treatments on small crop samples and starting with a low rate before full-scale implementation. The maximum cumulative amount of Sumagic applied must not exceed 10 ppm with coverage of 2 quarts per 100 sq. feet. This means that total amount used in sequential applications can only add up to 10 ppm spray (example, 1 application at 10 ppm or two applications at 5 ppm or 4 applications at 2.5ppm). The last spray must be no later than two weeks after the two to four leaf stage of development. Experiments have shown that sequential applications produce the best results and that the earlier that the plants receive the Sumagic spray, the greater effect it will have on the final height of the transplants. Since very few growth regulators are registered for vegetable bedding plants, plant height is often managed by adjusting temperature, water and fertilizer levels, or by physically brushing the plants. Research has shown that mechanical stress reduces stem elongation and maintains plant height. For example, brushing transplants twice daily for 18 days using about 40 strokes back and forth with a cardboard tube suspended from an irrigation boom can result in as much as a 30% reduction in stem elongation. Growers have also successfully used a wand made of plastic plumbing pipe or a flat piece of polystyrene foam. Vegetable plants such as tomatoes, eggplants and cucumbers have responded to this method of height control. Note that this technique has damaged some tender plant species such as peppers and could also enhance the spread of disease. The greater the difference between daytime and nighttime temperatures, the more plants will “stretch” (stems elongate).When the day temperature is very warm and the night temperature is cool or cold, plants will be taller. If the day and night temperature are both the same, plants will be shorter than with warm days and cool nights. If the night temperature in the greenhouse is kept warmer than the day temperature by using heating at night and ventilation during the day, the plants will be even shorter. Keeping day temperatures cool (70F) will help keep transplants shorter. The relationship is referred to as DIF, or difference between day minus night temperatures. Water stress is another tool growers can use to manage plant height. Maintaining plants on the dry side limits cell expansion and plant growth.

This method requires close attention to avoid permanent damage such as leaf burn or even plant death. One technique is to irrigate the growing mix thoroughly and then allow it to dry to the point where plants wilt before irrigating thoroughly again. Growth is restricted during the period when the growing medium is very dry. Once watered, the plants rapidly resume growth. Experienced tomato growers have successfully used this technique. Withholding nutrients can also be used to prevent stretching. Low phosphorus fertilization is especially effective for tomatoes. If carefully managed, a mild to moderate phosphorus (P) deficiency may result in a desirable reduction in growth with no foliar symptoms of P deficiency. See section on fertility for more information. Note: For field production the goal will be to put a vegetative plant out and promote rapid vegetative growth that will produce the largest yields. Water and nutrients must be carefully managed to produce healthy transplants.

ACCLIMATING OR HARDENING OFF TRANSPLANTS The transition from the greenhouse to the field involves changes in light, temperature and wind. Vegetable transplants benefit by a gradual “hardening” off period before they are transplanted into the field. Gradual exposure to outdoor growing conditions and reduced watering at the end of the growing period with some protection from wind and temperature but full exposure to light can increase the survival rate of transplants in the field. Three to six days are adequate to acclimate transplants. Care must be taken to not “overharden” young transplants. Cool-season crops exposed to very low temperatures can result in bolting (in cabbage) or buttoning (in broccoli or cauliflower). Warm-season crops generally are hardened at temperatures higher than those of cool-season crops. Cold temperatures can set back warm-season crops and can induce disorders such as catfacing in tomatoes.

INTEGRATED PEST MANAGEMENT There are only a limited number of insecticides and fungicides labeled

for greenhouse-grown vegetable bedding plants. Integrated pest management (IPM) offers a practical way to effectively manage pests on vegetable bedding plants and transplants. Through the use of sound cultural practices, monitoring techniques, accurate problem identification, and timely implementation and evaluation of appropriate management strategies, growers can improve their production while minimizing their reliance on routine pesticide applications. IPM utilizes many different management options; genetic, cultural, physical, mechanical, biological and chemical. Routine crop inspection alerts growers to developing pest and cultural problems while they are still minor and can be easily managed. Early detection and intervention is the foundation of an IPM program. Disease Management Diseases of vegetable bedding plants include Botrytis blight, dampingoff, Alternaria blight, Botrytis blight, late blight, powdery mildew, downy mildew, bacterial diseases such as bacterial leaf spot, bacterial canker, and black rot, and viral diseases such as Cucumber Mosaic Virus (CMV), Tobacco Mosaic Virus (TMV), and Tospoviruses. Effective control of diseases requires accurate identification. Failure of disease control is often because the cause was not accurately identified. Symptoms caused by poor cultural practices can also mimic disease symptoms. Fungicides cannot correct problems caused by high soluble salts, poor aeration or a nutrient imbalance. An integrated approach to disease management involves the use of resistant cultivars, sanitation, sound cultural practices and the proper use of the correct pesticide.

RESISTANT CULTIVARS Seed catalogues often feature disease resistant and tolerant varieties of vegetables. Utilize resistant varieties where feasible, but take some time to research the diseases that are giving you the most trouble to find other strategies to incorporate into the disease management plan.

SEED TREATMENTS FOR DISEASE MANAGEMENT Seed treatments are useful for many vegetable crops to prevent root diseases, as well as certain diseases carried on or within the seed. There are

two general types of seed treatment: eradicative and protective. Eradicative seed treatments use hot water or chlorine to kill disease-causing agents on or within the seed. They are useful in controlling certain seed-borne bacterial diseases such as bacterial leaf spot on pepper and tomato and bacterial canker on tomato. Protective seed treatments use fungicides on the seed surface to protect the seed against decay and soil-borne organisms such as damping off caused by Pythium, Phytophthora and Rhzoctonia. For more information regarding seed treatments, contact your seed sales representative, Extension vegetable specialist or plant pathologist. Sanitation Pest management on vegetable bedding plants and transplants begins with a clean, weed-free, disinfected greenhouse. Before growing the crop, the greenhouse should be cleared of plant debris, weeds, flats and tools. Empty benches, potting tables, storage shelves, tools and cell packs should be washed and disinfected with a sanitizing agent. It is important to thoroughly clean or power wash to remove organic debris from plastic containers before using a sanitizing agent. Bits of organic debris can be difficult to remove and the organic matter can be a source of disease causing pathogens if the plug trays are reused. After the greenhouse has been sanitized, care must be taken to avoid recontamination with pathogens. Purchase certified, disease-free seed from reliable sources. If possible, purchase seed that has been disinfested by chemical and/or heat treatment by the seed company. Potting media is easily re-infested by dirty hose nozzles or tools and unsanitary growing conditions. The floor of the greenhouse is a good source for diseases. Use a hook to keep the hose nozzles off the floor. Grow transplants off the ground in a wellventilated greenhouse. To prevent root rot diseases, avoid over-watering and over-fertilizing. Water early in the day to allow foliage to dry quickly to help prevent foliar diseases. Use separate greenhouses for vegetable seedlings and ornamental bedding plants. Separate greenhouses: 1) will protect vegetable seedlings from any insect pests that may migrate from ornamentals and plants that are held over; 2) will protect vegetable seedlings from Tospoviruses; 3) protect curcurbit seedlings from powdery mildew originating on verbena and 4) facilitate treatment of the vegetable seedlings if pesticides are needed.

Keep tomato transplant production separate from greenhouse tomato fruit production. Greenhouses with both young transplants and mature plants increase the risk of perpetuating diseases.

TECHNIQUES TO REDUCE HIGH HUMIDITY High relative humidity is one of the major contributing factors to Botrytis blight and powdery mildew, common fungal diseases of bedding plants. Warm air holds more moisture than cool air. During warm days, the greenhouse air is more humid. As the air cools in the evening, the moistureholding capacity drops until the dew point is reached. Water then begins to condense on surfaces. Humidity can be reduced by exhausting the moist air and replacing it with cooler outside air that is drier. The method and time it takes to heat and vent depend upon the heating and ventilation system in the greenhouse. In greenhouses with vents, turn the heat on and crack the vents open about one inch. The moist humid air escapes from the vents. In greenhouses with fans, activate the exhaust fans for a few minutes and then heat the greenhouse to raise the air temperature. Then, shut off the fans. A clock can be set to activate the fans. The cooler, outside air will lower humidity levels as it is warmed in the greenhouse. A relay may be needed to lock out the furnace or boiler until the fan shuts off so that flue gases are not drawn back into the greenhouse. This will also help to prevent air pollution damage (ethylene or sulfur dioxide) to sensitive seedlings. Heat and vent two or three times per hour in the evening after the sun goes down and early in the morning at sunrise. Heating and venting can be effective even if it is cool and raining outside. Air movement, even in a closed greenhouse, helps reduce moisture on the plant surfaces and surrounding the plants. Using horizontal airflow (HAF) can also reduce condensation. HAF fans keep the air moving in the greenhouse, helping to minimize temperature differentials and cold spots where condensation occurs. Air that is moving is continually mixed. The mixed air along the surface does not cool below the dewpoint so does not condense on plant surfaces. In addition, cultural practices can be used to reduce humidity within the plant canopy. These include proper watering practices and spacing of plants. Since most vegetable bedding plants are grown in flats that are spaced flat to

flat, reducing humidity within the canopy is difficult. Proper planting dates, plant nutrition, watering practices and height management techniques help to prevent lush, overgrown plants thereby reducing humidity within the canopy. Always water in the morning to reduce the length of time the leaves stay wet after irrigating to prevent foliar diseases. Rising temperatures during the day will evaporate water from the foliage, so the leaves stay dry. Avoid watering late in the day or when water will sit on leaf surfaces for long periods of time. Fungicides Fungicides can provide excellent management of some diseases, but for others they may be ineffective. In general, to control root diseases, broadspectrum fungicides should be applied as a drench on a preventative basis. Read directions for application on pesticide labels. An application of additional water may be necessary. For foliage diseases, obtain thorough spray coverage and treat when disease is first evident. provides a listing of fungicides labeled for vegetable bedding plants. Biofungicides Biofungicides are biological fungicides that contain living organisms such as fungi, bacteria, or Actinomycetes (a group of bacteria that form branching filaments) that attack plant pathogens and the diseases they cause. They can be used as part of an integrated disease management program to reduce the risk of pathogens developing resistance to traditional fungicides. Currently, there are no pathogens that are resistant to biological fungicides. Biological fungicides may suppress diseases in a number of different ways. They may directly compete with the pathogen. The biological fungicide “shields” the roots by growing a defensive barrier around the roots. The microorganisms may produce an antibiotic or another toxin that kills the target organism. They may attack and feed upon the pathogen (mycoparasitism). As such, the biological fungicide must be present at the same time or before the pathogen appears. Some biological fungicides induce the plant to turn on their own defense mechanisms. Some of the advantages of using biological fungicides include: lower re-entry interval (REI) than many traditional fungicides, may be organic products (OMRI listed), may be less phytotoxic, and many can be used in rotation with synthetic chemicals.

(See company web sites for more information on compatibility). Biofungicides should be used as a preventative treatment in conjunction with a regular monitoring program where root health and crop quality is evaluated. They will not cure diseased plants and must be applied before the onset of the disease. Biological fungicides need to be used in conjunction with standard cultural practices to help prevent diseases. Storage conditions, soil and air temperatures, and use of other chemicals affect their efficacy. Most biological fungicides also have a limited shelf life of one year. A number of products are commercially available for use on vegetable bedding plants and transplants, please refer to the label for specific information regarding what specific types of diseases and specific plants are listed for each individual biological fungicide.

SPECIFIC DISEASES Blights Botrytis blight Botrytis can cause leaf blight, stem cankers, damping off and root rot. Plants may be attacked at any stage, but the new tender growth, freshly injured tissues and dead tissues are most susceptible. Symptoms Botrytis blight produces characteristic gray fuzzy appearing spores on the surface of infected tissues. Tan stem cankers can develop on basil. Air currents and splashing water can easily disseminate the spores. In general, germination of spores and infection is dependent on a film of moisture for 8 to 12 hours, relative humidity of 93% or greater and temperatures between 55° and 65°F. After infection, colonization of plant tissues can occur at temperatures up to 70°F. Management Botrytis diseases can only be managed by a combination of methods including manipulation of environmental conditions (temperature, humidity and duration of leaf wetness), sound cultural practices and use of fungicides. Fungicides alone cannot control Botrytis and this pathogen has a long history

of fungicide resistance development • Control weeds and remove plant debris before and during production. • Dispose of diseased plants and debris in a plastic trash bag. Keep the bag closed to help prevent spreading spores to uninfected plants as the bag is removed from the greenhouse. Cover trash cans to prevent the airborne spread of spores from diseased plant tissue. • Reduce humidity and leaf wetness duration to prevent spore germination. Provide good air circulation and reduce humidity within the canopy. • Proper planting dates, fertility, watering and height management will prevent overgrown plants, thereby reducing humidity within the canopy. • Water in the morning, never late in the day. Rising temperatures during the day will cause water to evaporate from the foliage and dry the leaf surface. • Avoid growing ornamental hanging baskets above vegetable bedding plants. Spent flowers dropping on plants below causeBotrytis infection. Late Blight

Late Blight

Late blight is caused by the water mold Phytophthora infestans. The fungus typically overwinters in potato cull piles or in soil where plant tissue has not completely frozen and is not considered a problem for locally grown tomato seedlings. Symptoms Found on tomato and potato plants, late blight is caused by the fungus Phytophthora infestans and is common throughout the United States. True to its name, the disease occurs later in the growing season with

symptoms often not appearing until after blossom. Late blight first appears on the lower, older leaves as water-soaked, graygreen spots. As the disease matures, these spots darken and a white fungal growth forms on the undersides. Eventually the entire plant will become infected. Crops can be severely damaged. Unlike other fungal diseases, this plant problem does not overwinter in the soil or on garden trash. Instead the spores are introduced by infected tubers, transplants or seeds. Wind will also carry the disease from nearby gardens. Warm temperatures (70-80°F) and wet, humid conditions promote its rapid spread. Management Many of the same fungicides used for Botrytis blight will help protect tomato seedlings from late blight. Damping-off of Seedlings Damping-off is a common disease of germinating seeds and young seedlings. Several fungi are capable of causing damping-off including Rhizoctonia, Alternaria, Sclerotinia and the water molds, Phytophthora and Pythium. Soil-borne fungi generally do not produce air-borne spores but are easily transported from contaminated soil to pathogen-free soil by infected tools, hose ends, water-splash and hands. Young seedlings are most susceptible to damping-off. However, later in the crop cycle, the same pathogens may cause root and stem rot. Symptoms A soil-borne fungal disease that affects seeds and new seedlings, damping off usually refers to the rotting of stem and root tissues at and below the soil surface. In most cases, infected plants will germinate and come up fine, but within a few days they become water-soaked and mushy, fall over at the base, and die.

Damping-off Seedlings

Several fungi can cause decay of seeds and seedlings, including species of Rhizoctonia, Fusarium and Phyto-phthora. However, species of the soil fungus Pythium are most often the culprit. Damping off typically occurs when old seed is planted in cold, wet soil and is further increased by poor soil drainage. High humidity levels, rich potting soils and planting too deeply will also encourage its growth. Fungal spores live in the soil and are primarily a problem in seed beds. They can be transported on garden tools and in garden soils taken into the house or greenhouse. .Management Damping-off must be prevented because it is difficult to stop once symptoms occur. There are several strategies to prevent damping-off. • Use only certified disease-free seed from reputable seed companies. • Use fungicide-treated seed. Certain fungicides are labeled for dampingoff for selected vegetable crops. • Use pasteurized soil, properly produced compost-based or soilless mixes. Incorporate biological fungicides into your soilless mix or apply biological fungicides as a drench at planting. • Disinfect all flats, cold frames, pots and tools. • Germinate seed under conditions that will ensure rapid emergence, such as with the use of bottom heat. • Avoid overwatering, excessive fertilizer, overcrowding, poor air circulation, careless handling, and planting too deeply. • Fill flats with pre-moistened growing media to avoid compaction.

Lightly fill and brush containers. Do not pack young plants into containers, use pre-dibbled holes for transplants. • To avoid compaction, do not stack or “nest” filled trays or pots. • Provide adequate light for rapid growth. • Discard entire infected flats. Downy Mildew

Downy Mildew

Although they have traditionally beenincluded taxonomically with true fungi,these organisms and their relatives in thegenera Phytophthora and Pythium andothers in the Oomycota are now notbelieved to be closely related to true fungi.Morphologically, they are similar to fungiand have absorptive nutrition. Thechemicals used to control downy mildewsare similar to those used for Pythium andPhytophthora and different from most of those used for true fungi. Symptoms A fungal disease, downy mildew (Plasmopara viticola) affects many plants and appears as yellow to white patches on the upper surfaces of older leaves. On the undersides, these areas are covered with white to grayish, cotton-like fungi. These “downy” masses are most often noticed after rain or heavy dew and disappear soon after sunny weather resumes. As the disease progresses leaves may eventually turn crisp and brown and fall off even though the plant has ample water. Downy mildew occurs mostly in cool, moist weather, usually in early spring or late fall. Spore production is favored by temperatures cooler than 65ÚF. and by relative humidities approaching 100%. This disease overwinters on plant debris and in the soil. Fungal spores can be carried by

insects, wind, rain or garden tools. Management Management of environmental conditions such as temperature, humidity and duration of leaf wetness, sound cultural practices and fungicides will help prevent disease development. • It is vital to reduce humidity and leaf wetness duration to prevent spore germination. See techniques for reducing relative humidity. • Provide good air circulation and reduce humidity within the canopy. Proper planting dates, fertility, watering and height management will prevent overgrown plants, thereby reducing humidity within the canopy. • Water in the morning, never late in the day. Rising temperatures during the day will cause water to evaporate from the foliage and dry the leaf surface. Powdery Mildew Powdery mildew is a common disease on many types of plants. There are many different species of powdery mildew fungi (e.g., Erysiphe spp., Sphaerotheca spp.) and each species only attacks specific plants. A wide variety of vegetable crops are affected by powdery mildews, including artichoke, beans, beets, carrot, cucumber, eggplant, lettuce, melons, parsnips, peas, peppers, pumpkins, radicchio, radishes, squash, tomatillo, tomatoes, and turnips. Powdery mildews generally do not require moist conditions to establish and grow, and normally do well under warm conditions; thus they are more prevalent than many other leaf-infecting diseases under California’s dry summer conditions.

Powdery Mildew

Symptoms Common on many plants and easily recognized, powdery mildew is a fungal disease found throughout the United States. It is caused by a variety of closely related fungal species, each with a limited host range. (The fungi attacking your roses are unlikely to spread to your lilacs). Low soil moisture combined with high humidity levels at the plant surface favors this disease.

Brownish spots on pea pod from powdery mildew infection.

Symptoms usually appear later in the growing season on outdoor plants. Powdery mildew starts on young leaves as raised blister-like areas that cause leaves to curl, exposing the lower leaf surface. Infected leaves become covered with a white to gray powdery growth, usually on the upper surface; unopened flower buds may be white with mildew and may never open. Leaves of severely infected plants turn brown and drop. The disease prefers young, succulent growth; mature leaves are usually not affected.

Fungal spores overwinter inside leaf buds and other plant debris. Wind, water and insects transmit the spores to other nearby plants. Zucchini, roses and zinnia are especially susceptible. Management The best method of control is prevention. Planting resistant vegetable varieties when available, or avoiding the most susceptible varieties, planting in the full sun, and following good cultural practices will adequately control powdery mildew in many cases. However, very susceptible vegetables such as cucurbits (cucumber, melons, squash, and pumpkins) may require fungicide treatment. Several least-toxic fungicides are available but must be applied no later than the first sign of disease. Plant in sunny areas as much as possible, provide good air circulation, and avoid applying excess fertilizer. A good alternative is to use a slowrelease fertilizer. Overhead sprinkling may help reduce powdery mildew because spores are washed off the plant. However, overhead sprinklers are not usually recommended as a control method in vegetables because their use may contribute to other pest problems. Fungicide Application In some situations, especially in the production of susceptible cucurbits, fungicides may be needed. Fungicides function as protectants, eradicants, or both. A protectant fungicide prevents new infections from occurring whereas an eradicant can kill an existing infection. Apply protectant fungicides to highly susceptible plants before the disease appears. Use eradicants at the earliest signs of the disease. Once mildew growth is extensive, control with any fungicide becomes more difficult. The products listed here are for home garden use. Fungicides Several least-toxic fungicides are available, including horticultural oils, neem oil, jojoba oil, sulfur, and the biological fungicide Serenade. With the exception of the oils, these materials are primarily preventive. Oils work best as eradicants but also have some protectant activity. Oils To eradicate mild to moderate powdery mildew infections, use a

horticultural oil such as Saf-T-Side Spray Oil, Sunspray Ultra-Fine Spray Oil, or one of the plant-based oils such as neem oil or jojoba oil (e.g., E-rase). Be careful, however, to never apply an oil spray within 2 weeks of a sulfur spray or plants may be injured. Also, oils should never be applied when temperatures are above 90°F or to drought-stressed plants. Some plants may be more sensitive than others, however, and the interval required between sulfur and oil sprays may be even longer; always consult the fungicide label for any special precautions. Sulfur Sulfur products have been used to manage powdery mildew for centuries but are only effective when applied before disease symptoms appear. The best sulfur products to use for powdery mildew control in gardens are wettable sulfurs that are specially formulated with surfactants similar to those in dishwashing detergent (e.g., Safer Garden Fungicide) However, sulfur can be damaging to some squash and melon varieties. To avoid injuring any plant, do not apply sulfur when air temperature is near or over 90°F and do not apply it within 2 weeks of an oil spray. Other sulfur products, such as sulfur dust, are much more difficult to use, irritating to skin and eyes, and limited in terms of the plants they can safely be used on. Copper is also available to control powdery mildew but is not very effective. Biological Fungicides Biological fungicides (such as Serenade) are commercially available beneficial microorganisms formulated into a product that, when sprayed on the plant, destroys fungal pathogens. The active ingredient in Serenade is a bacterium, Bacillus subtilis, that helps prevent the powdery mildew from infecting the plant. While this product functions to kill the powdery mildew organism and is nontoxic to people, pets, and beneficial insects, it has not proven to be as effective as the oils or sulfur in controlling this disease. Bacterial Diseases Bacterial diseases of vegetable bedding plants, such as bacterial leaf spot of peppers and tomatoes, bacterial speck & bacterial canker of tomatoes, and black rot on cole crops are introduced into a greenhouse through infected seed and transplants. Bacterial leaf spot, Bacterial speck

Xanthomonas campestris

Bacterial spot is one of the most devastating diseases of pepper and tomato grown in warm, moist environments. Once present in the crop, it is almost impossible to control the disease and prevent major fruit loss when environmental conditions remain favorable. The study of this bacterial pathogen also has significantly enhanced the scientific understanding of hostpathogen interactions and the molecular basis of the gene-for-gene model. Bacterial spot is caused by several species of gram-negative bacteria in the genus Xanthomonas. In culture, these bacteria produce yellow, mucoid colonies. A “mass” of bacteria can be observed oozing from a lesion by making a cross-sectional cut through a leaf lesion, placing the tissue in a droplet of water, placing a cover-slip over the sample, and examining it with a microscope (~200X)

Bacterial spot lesions on pepper fruit and the peduncle

Symptoms Bacterial leaf spot is caused by Xanthomonas campestris pv. vesicatoria and is found primarily on peppers although all aboveground parts of tomatoes are also susceptible. Spots on leaves are chocolate-brown with yellowing at lesion’s margins and irregularly shaped with areas of dead leaf tissue. At first, the spots are less than ½ of an inch in diameter. Severely spotted leaves

will appear scorched and defoliation may occur. This disease is most prevalent during moderately high temperatures and long periods of leaf wetness. Bacterial speck occurs on tomato but not pepper. The bacterium, Pseudomonas syringae pv. tomato, causes small black spots to develop resulting in chlorosis (yellowing), necrosis (dead tissue) and blighting of the foliage. Bacterial speck can usually be distinguished from bacterial spot by the size of the lesions, however, in some cases, the symptoms look similar. Bacterial canker

Bacterial canker

Bacterial canker of tomato is caused by Clavibacter michoiganensis pv. michiganensis (formerly Corynebacterium michiganense).In New England, bacterial canker occurs less frequently than other tomato diseases but it is potentially more destructive. The bacterium is seed-borne but may survive on plant debris in soil for at least one year. It can also survive in the greenhouse on wooden stakes and flats. Wilt, leaf scorch, canker, pith necrosis and fruit spot may occur singly or in combination depending on the circumstances. When the bacterium is carried in the seed, the vascular system becomes colonized, resulting in wilt, pith necrosis and external cankers. Wilt initially occurs on one side of a leaf or one half of a plant because only a portion of the vascular system is blocked. Cankers and pith necrosis occur in later stages of disease development. Cankers are dark and water-soaked in appearance and often exude bacteria that are easily spread to adjacent plants. Pith necrosis is first evident as a darkening of the center of the stem that soon becomes chambered or hollow. When leaf scorch occurs, the petioles usually bend downward while the leaf edges curl up. The margins of the leaves become brown with a yellow border to the inside. Scorching of the foliage

often develops in the absence of wilt or stem canker. Transplants may not express symptoms until six to eight weeks after infection and initial symptom expression is accelerated by environmental stress. Symptoms Bacterial canker is most common on cherries and plums, but may also affect apricots, peaches and many other kinds of stone fruits. Suspect this plant disease if sunken, water-soaked or “gummy” lesions form on the trunk or twigs. When trees begin active growth in the spring, a sour smelling sap may ooze from these wounded areas. The cankers become darker than the surrounding healthy bark, and the underlying tissue is reddish-brown to black and moist. Black rot Black rot, caused by the bacterium Xanthomonas campestris pv. campestris occurs where cruciferous plants are grown. All Brassicas can be severely affected. The bacterium enters the leaves by colonizing the hydathodes (water pores) and moves from the leaf margins inward. Lesions may also begin at wounds. Diseased tissue is often V-shaped; flaccid, tan to yellow and with blackened veins. The blackened veins are diagnostic and are best seen by holding the leaf up to the light. When the lesions reach the petiole and stem, the bacterium moves systemically through the plant, resulting in premature leaf drop. At this stage of disease, a cross-section of the stem will reveal a ring of discolored vascular tissue. Symptoms In the field, the disease is easily recognized by the presence of large yellow to yellow-orange “V”-shaped areas extending inward from the margin of a leaf, and by black veins in the infected area. Usually only a few of the outer leaves are involved. If infection occurred in a young seedling, the disease is usually much more severe since the main stem becomes infected and the disease becomes systemic in the plant. These plants remain stunted and the veins in the stems are black. The heads from these plants deteriorate rapidly after harvest. Although the distribution of diseased plants in the field may be quite uniform, the disease may be more common and severe in low and shaded areas. If a few infected seedlings were set in the field, scattered pockets of

diseased plants will appear in the field early in the growing season. Diseased plants often appear in the same rows as a result of spread during cultural operations. Seedling infection is often very difficult to detect. Infected seedlings tend to be stunted and often exhibit one-sided growth. The leaves may be light green, and lower leaves may drop prematurely. Vascular elements in the stems will be black. However, infected seedlings may show no symptoms at all. The recognition of seedling infection is made more difficult since only a few (less than 1%) of the seedlings in a lot may have the disease. The recognition of infected seedlings is very important in the control of the disease. Management of bacterial diseases These bacteria can be introduced on infected seeds, infected transplants purchased from another operation, or in the field on crop residues. These bacteria can also survive on weeds in the same family as the host crop. The management of these bacterial diseases is similar and includes the following strategies: • Buy certified disease-free seed from a reputable source. • Use hot water-treated seed. Ideally, the seed should be custom-treated by the seed company. Seed companies may treat the seed upon request. There is a risk that germination percentages will be reduced if the seed crop is grown under stressful environmental conditions. • Promptly remove infected plants and adjacent plants to prevent further infection and avoid unnecessary handling of plant material. • Avoid overhead irrigation, splashing or periods of extended leaf wetness. • Disinfect all benches, equipment, flats and stakes. • Follow sound practices for weed and insect control. • Prevent bacterial leaf spot on peppers by choosing resistant varieties whenever possible. There are many resistant varieties of bell peppers available, but few resistant specialty peppers. Viral Diseases Some viral diseases of vegetable bedding plants include cucumber

mosaic virus (CMV), tobacco mosaic virus (TMV) and the tospoviruses, INSV and TSWV. There is no control for plants infected with a virus. It is important to have the virus disease accurately identified. Serological techniques such as ELISA (enzyme-linked immunosorbent assay) are now available to accurately identify a wide range of viruses. On-site grower kits using this same technology are also available from Agdia to test for viruses such as CMV, TMV, INSV, and TSWV. Cucumber mosaic virus Cucumber mosaic virus (CMV) has a wide host range of over 400 species of plants including vegetables, ornamentals and weed hosts. Symptoms CMV infects 1200 species in over 100 plant families and can cause significant economic losses in many vegetable and horticultural crops. CMV causes a systemic infection in most host plants, but may remain symptomless in some crops like alfalfa. Symptoms of cucumber mosaic can vary greatly depending on the crop infected and the age of the plant when infection occurs. Cucurbits: Almost all cucurbits are susceptible to CMV, with symptoms varying in severity. Severe epinasty, downward bending of the petiole and leaf surface along with leaf reduction, are common in early season infection of summer squash. Plants infected early in the season are severely stunted and leaves are malformed, and fruit are unmarketable because of pronounced rugosity (roughness) on the fruit surface, as shown on the infected zucchini plant and fruit in Figure 3B. Infection of vining crops, such as muskmelon, show severely stunted growing tips, and although fruit may not show symptoms they are of poor quality. If the yellow squash variety grown lacks the precocious gene, color breaking will occur on the fruit, causing the fruit to show green blotchy patterns, but these symptoms are absent in yellow squash varieties with the precocious gene. Color breaking on fruit of varieties without the precocious gene will also occur with Watermelon mosaic potyvirus (WMV) infection; however this protection does not hold true for Papaya ringspot potyvirus or Zucchini yellow mosaic potyvirus, where both foliage and fruit of yellow squash are severely affected. Pumpkin is another

cucurbit, that when infected at any early stage, will express severe foliar mosaic and the fruit will show a mosaic pattern and would be unmarketable. Pepper: Foliar symptoms of pepper plants vary with stage of infection. The initial flush of systemic symptoms typically includes a chlorosis of young leaves that may occur over the basal portion of the leaf or over the entire leaf. Oak leaf and ringspot patterns may develop on these leaves as the plant ages. As new leaves emerge, these leaves develop a chlorotic mosaic pattern that tends to encompass the entire leaf. Leaves that develop subsequent to those expressing the chlorosis and chlorotic mosaic symptoms may have varied degrees of deformation including sunken interveinal lamina with protruding primary veins. These leaves also have a dull light green appearance as opposed to the dark green, rather shiny leaves of healthy pepper plants. These symptom patterns vary in severity depending on the age of the plant at the time of infection with more severe symptoms typically resulting when plants are young at the time of infection. CMV infected plants also tend to be stunted with those plants infected early in development possibly expressing severe stunting, whereas plants infected at later stages of development may have little, if any, stunting. Pepper fruit may develop ringspotting and roughness leading to unmarketable fruit.

Fig. 15.9 a-i. CMV symptoms on non-cucurbit crops a. Oak leaf and ringspot patterns on pepper. b. Malformed, dull light green leaves on pepper. c. Ringspotting and roughness of pepper fruit. d. Chlorosis of spinach. e. Severe roughness of the leaf and

occasional necrosis within the leaf tissue of Romaine lettuce. f. Yellowing and veinal necrosis of celery. g. Elongated, brown to translucent, sunken beige colored lesions on celery stalk. h. Filiformity or shoestring-like leaf blades of tomato. i. Leaf curl, green mottle and blistering of beans.

Spinach: CMV infection of spinach is often referred to as spinach blight. The symptoms can vary depending upon the variety, plant age when infected, temperature, and virus strain. Typical symptoms include leaf chlorosis, which can progress to cause severe blighting of the growing point and eventual plant death. In addition to chlorotic mottle, leaves can show narrowing, wrinkling with vein distortion, and inward leaf roll. Lettuce: Symptoms of CMV infection of lettuce consist of leaf mottling, severe roughness of the leaf and occasional necrosis within the leaf tissue. Plants are usually stunted if infected at an early stage of development. Celery: CMV infection of celery is referred to as southern mosaic virus in the older literature following its description in southern production areas. Initially leaves will develop veinclearing and mosaic, and later these same leaves may show yellowing and veinal necrosis. The petioles of these plants show elongated, brown to translucent, sunken beige colored lesions, making the celery stalk unmarketable. Symptoms may also be transient, such that later in the season the symptoms may be limited to a dull cast, while under cool growing conditions the obvious symptoms may be muted. Tomato: Tomato plants infected with CMV in the early stages are yellow, bushy and considerably stunted. The leaves may show a mottle similar to that caused by Tomato mosaic virus (ToMV). The most characteristic symptom of CMV is filiformity or shoestring-like leaf blades. Management: Rogue diseased plants. Eliminate weeds such as common pokeweed, chickweed, field bindweed, yellow rocket, and bittersweet nightshade that may be reservoirs of CMV. Genetic resistance for Cucumber mosaic virus (CMV) in melon (Cucumis melo) is derived from oriental melons, and depending on the source, resistance is controlled by two or three complementary recessive genes. Some of these factors for resistance are strain and/or temperature dependent, with plants developing symptoms at temperatures below 20°C. No commercial muskmelon varieties with CMV resistance are available. Fortunately, most varieties of watermelon (Citrullus lanatus) are resistant to the most prevalent strains of CMV, with the exception of a specific strain that

can infect plants systemically. CMV resistance in spinach is controlled by a single dominant gene in the variety Virginia Savoy, and this has been incorporated into many current spinach varieties. This resistance, however, is not complete and may break down at temperatures greater than 28°C. Virus resistance in lettuce to CMV was identified in Lactuca saligna PI 26153 from Portugal, a distantly related species of lettuce (L. sativa). However, this resistance is strain specific. An accession from L. serriola proved to be tolerant to three strains of CMV studied, but this tolerance has not been transferred to any current commercial variety. Extensive effort was deployed to develop pepper resistance for CMV, and although tolerance has been described in some varieties, such as ‘Perennial’, it was not utilized in any commercial variety. More recently the pepper variety ‘Peacework’ was developed with a high level of resistance. Sources of resistance and tolerance to CMV have been reported in wild tomato relatives but not in Solanum lycopersicum. No tomato varieties with CMV resistance have been released. Eradication of weed hosts is often a difficult task because of the extensive host range of this virus. However, elimination of several of the key perennial or biennial weeds located near the crop may reduce severe virus pressure, and has successfully been used to control CMV. Tobacco Mosaic Virus (TMV) TMV is made up of a piece of nucleic acid (ribonucleic acid; RNA) and a surrounding protein coat. The complete virus is a submicroscopic, rigid, rod-shaped particle. Once inside the plant cell, the protein coat falls away and nucleic acid portion directs the plant cell to produce more virus nucleic acid and virus protein, disrupting the normal activity of the cell. TMV can multiply only inside a living cell but it can survive in a dormant state in dead tissue, retaining its ability to infect growing plants for years after the infected plant part died. Most other viruses die when the plant tissue dies. The most important way that TMV can be spread from plant to plant is on workers’ hands, clothing or on tools. This is called ‘mechanical’ transmission. When plants are handled, the tiny leaf hairs and some of the outer cells inevitably are damaged slightly and leak sap onto tools, hands, and

clothing. If the sap contains TMV, it can be introduced into other plants when those come in contact with this sap. Sucking insects such as aphids do not spread TMV. Chewing insects such as grasshoppers and caterpillars occasionally spread the virus but are usually not important in spread. Vegetative propagation perpetuates TMV and other virus diseases. Cuttings taken from an infected plant usually are infected even if no symptoms are immediately exhibited by the cutting. The virus particles are found in all parts of the plant except the few cells at the tips of the growing points. Infected stock plants should be discarded immediately. TMV can also survive outside the plant in sap that has dried on tools and other surfaces. If a TMV plant is handled and then you open a door with that hand, you have now put TMV on the door handle. The next person to open the door can pick up the TMV and spread it to any plant that they touch. Tobacco products, particularly those containing air-cured tobacco, may carry TMV. Flue-cured tobacco, used in making cigarettes, is heated repeatedly during its processing, thereby inactivating most if not all TMV. When tobacco products are handled or kept in pockets, hands and clothing can become contaminated with TMV and be a source of virus. TMV is NOT spread in the smoke of burning tobacco. Symptoms Symptoms include yellow mottling, upward leaf curling and overall stunting. Some infected plants may not show any symptoms at all. Management Discard infected plants including roots. Disinfect hands by washing with milk, or tri-sodium phosphate and then thoroughly with soap and water. Smokers need to wash their hands before entering the greenhouse so they do not infect plants. In greenhouses, hard surfaces such as doorknobs, or flats can become contaminated after handling virus-infected plants and remain a source of infection. Thoroughly disinfect the growing area with a commercial disinfectant. Control perennial weeds in the solanaceous family such as ground cherry and horsenettle that could be reservoirs of TMV. Tospoviruses Tospoviruses are a group of viruses that include impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV). They may infect

hundreds of plant species including basil, tomatoes, peppers and eggplant. These viruses are primarily spread by the western flower thrips. Tospoviruses are not seedborne but are brought into the greenhouse on vegetatively propagated ornamental plants or seedlings that have been exposed to the virus. Once the thrips in the greenhouse become infected, they can transmit the virus to susceptible crops and weeds. Symptoms Symptoms include stunting, foliar ringspots and black lesions on stems. Symptoms of INSV and TSWV will vary depending upon the host. Management To manage Tospoviruses, it is necessary to discard infected plant material, including weeds and to manage thrips. Infected vegetable transplants planted into the garden or field will be stunted and will not produce a harvestable crop. Since INSV and TSWV are not seed-borne, vegetable transplants may be kept free of Tospoviruses if they are not brought into contact with other infested crops or thrips carrying the virus. Growers attempting to concentrate all their warm temperature crops in a single house run a risk of mixing Tospovirus-free vegetable seed crops with leftover ornamental stock plants or new cuttings that may carry the virus. Prefinished or vegetatively propagated ornamentals from another producer could be infested with thrips or a virus. Therefore, vegetable bedding plants should always be grown in separate greenhouses.

GENERAL INSECT AND MITE PEST MANAGEMENT Monitoring A regular monitoring program is the basis of all pest management programs. Conduct a regular, weekly scouting program to detect pest problems early. This early detection and treatment will result in better pest control since plant canopies are smaller and better spray coverage can be achieved. Yellow Sticky Cards Use yellow sticky cards to trap and detect adult stages of flying insects such as western flower thrips (WFT), whiteflies, fungus gnats, shoreflies,

leafminers and winged aphids. Remember that mites and wingless aphids do not fly and will not be caught on the cards. Place one to four cards per 1,000 square feet. The cards should be spaced equally throughout the greenhouse in a grid pattern with additional cards located near doorways and vents. Place some cards just above the plant canopy (to detect thrips and whiteflies) and some of the cards on the rim of the flats or pots to detect fungus gnats. Inspect and replace the cards weekly to keep track of population trends. Plant Inspection Plant inspection is needed to assess general plant health and to detect diseases, mites and wingless aphids plus any hot spots of immature whiteflies. Randomly select plants at ten locations in an area of 1,000 square feet, examining plants on each side of the aisle. Start this pattern at a slightly different location each week, walking through the greenhouse in a zigzag pattern down the walkway. Examine the underside of leaves for insect pests and inspect root systems to determine whether they are healthy. Key Plants and Indicator Plants Focus on scouting key plants and indicator plants. Key plants are those plants or cultivars that have serious, persistent problems every year. For example, peppers and eggplants are prone to aphid infestations. Look for aphids on the young leaves and for shiny honeydew on the upper leaf surface. If grown near flowering plants, peppers and eggplant will also indicate an early thrips population. Look for distorted, young leaves with silvery flecked scars that are signs of thrips feeding damage. Fava beans and certain cultivars of petunia are used as indicator plants to detect the presence of thrips carrying INSV and TSWV. These plants will develop viral symptoms within one week if fed on by the infected thrips. The petunia cultivar ‘Summer Madness’ and several varieties of fava bean have been successfully used to detect tospoviruses. To use petunias and fava beans as indicator plants: •

Remove flowers from indicator plants to encourage feeding on foliage where symptoms can be observed. • Place a blue non-sticky card in each pot at plant height. The blue card will attract thrips to the indicator plant. Blue plastic picnic plates work well.

• Place petunia plants throughout the greenhouse among the crop at a rate of one plant every 20-30 feet and fava bean plants at the rate of 12 pots per 1,000 sq. ft. • Remove symptomatic leaves on petunia plants and continue to use the plants. The virus is not systemic in these plants. Thrips feeding injury leaves distinct white feeding scars on the foliage. Virus symptoms appear as a brown rim around the feeding scars. • Remove entire plants of fava beans if symptoms are observed, because the virus is systemic in these plants. Viral symptoms appear as dark brown angular lesions on leaves or yellow to light green ring spots. Dark necrotic areas can also be seen on the stem. Fava beans have dark black spots on their stipules that should not be confused with viral symptoms. • Replace with new plants, planting 1-2 bean seeds per 4’’ or 6 “pot.

RECORD KEEPING AND DECISION-MAKING Each time the crop is scouted, record the pest numbers, their location and the number of plants inspected. Records on pest numbers and locations will help you identify population trends. Population trends will also indicate if initial control measures were successful or if they need to be repeated. Once this information is collected each week, a pest management decision can then be made. Monitoring and record keeping will answer the following questions and help you make the necessary treatment decisions. Is the population decreasing, increasing or remaining stable over the growing season? Do you need to spray? Are insects and mites migrating from weeds under the benches to your crops? Is the treatment from last week working?

BIOLOGICAL CONTROL FOR INSECTS AND MITES Biological control may be an option for aphids, mites, fungus gnats, thrips and whiteflies. Natural enemies are living organisms that need to be released when pest populations are low. They do not act as quickly as pesticides so cannot be used as a “rescue” treatment. Natural enemies (parasites, predators or pathogens) are best used early in the cropping cycle when plants are small, pest numbers are low and damage is not yet observed. A detailed plan of action is needed to insure success. Accurately identify the

key pests in your production system. Natural enemies, especially parasites, are often very specific to a particular pest. Many insecticide residues can adversely affect natural enemies for up to 3 months after their application. A separate greenhouse is best. With help from your biological control agent supplier, establish a schedule for introducing the natural enemies. Release rates and timing will vary depending upon the crop and its size, the degree of infestation, effectiveness and type of natural enemies, plus the time of year. Starting a biological control program will involve some trial and error, as release rates have not been scientifically evaluated for vegetable bedding plants. Vegetable bedding plants with only one or two key insect pests or with a longer production schedule may be logical candidates for biological control. Be sure that natural enemies are received from your supplier quickly (4 days), and that they are not exposed to temperature extremes (too hot or cool) during shipment. Request that the biological control supplier include ice packs and a data logger (if possible). Inspect natural enemies for viability and quality when they are received.

SPECIFIC INSECT PESTS AND MITES

Common insect pests on vegetable bedding plants include aphids, fungus gnats, shore flies, whiteflies, thrips and two-spotted spider mites. The following are brief descriptions, life cycles and monitoring tips for the major pests. Aphids Lifecycle Several species of aphids can occur on vegetable transplants, but the

most common are green peach, melon and foxglove. Aphids are small, 1/16inch in length, round, soft-bodied insects that vary in color from light green to pink or black. The green peach aphid is yellowish-green in summer; pink or yellowish in fall and spring. Winged forms are brown with a large dusky blotch on the abdomen. Melon aphids are greenish-yellow to very dark green with black mottling and short dark cornicles or “tailpipes” (tubular structures on the posterior part of the abdomen). Foxglove aphids are smaller than potato aphids but larger than melon and green peach aphids. The foxglove aphid is a shiny light yellowish green to dark green in color with a pear-shaped body. The only markings on the bodies of wingless adults are dark green patches at the base of the cornicle. The legs and antennae also have black markings. Foxglove aphids cause more leaf distortion than green peach or melon aphids. Aphids feed by inserting their piercing, sucking mouthparts into plant tissue and removing fluids. In greenhouses, aphids are usually females that produce live young called nymphs. Each female can produce 50 or more nymphs. Nymphs mature to adulthood and begin reproducing in as little as 7 to 10 days. Adults are usually wingless, but some will produce wings when populations reach outbreak levels. Large numbers of aphids will stunt and deform plants. In addition, aphids produce a sticky digestive by-product called honeydew and their white shed cast skins may be unsightly. Sometimes, these white cast skins are mistakenly identified as whiteflies. Honeydew can cover leaves and provide a food source for a superficial black fungus known as sooty mold. Aphids are present on weeds and winged aphids may enter the greenhouse through vents. Aphids can also transmit certain viruses. Monitoring Examine the foliage, along stems and new growth of key plants such as peppers, eggplants, cole crops and leafy greens to detect an early aphid infestation. Signs of aphid activity include shed white skins, shiny honeydew, curled new leaves, distorted growth and the presence of ants. Yellow sticky cards help detect the entrance of winged aphids into the greenhouse from outdoors. Yellow cards will not, however, allow you to monitor aphids within the crop, as most of the aphids will be wingless. Whiteflies

Greenhouse whitefly adults

The major foliar pest in greenhouses is the silverleaf whitefly, Bemisia argentifolii. This insect first became a greenhouse pest in the U.S. in 1986 when it became more common than the greenhouse whitefly (Trialeurodes vaporariorum), the primary whitefly pest of poinsettia at that time. The current problem with B. argentifolii began in 1986 when poinsettia growers experienced devastating outbreaks of what was at first called sweetpotato whitefly, Bemisia tabaci. Sweetpotato whitefly had been in the U.S. since 1897 but was not a serious agricultural pest. To distinguish this new, damaging form of sweet potato whitefly, from the pre-existing, less damaging strain, the old strain was referred to as strain A, and the new strain damaging poinsettia, field crops, and vegetables, was called strain B. B. tabaci strain B had spread to almost every poinsettia growing region in the United States. Work on the biology, morphology, and genetics of strains A and B revealed sufficient difference that strain B was described as a new species of whitefly. Strain B was named Bemisia argentifolii, the silverleaf whitefly, because of its ability to cause squash silverleaf. The exact country of origin of B. argentifolii is unknown, but it is thought to be India or Pakistan as this area shows the greatest diversity in parasitoids of Bemisia, which may be an indication of a genus epicenter. Whiteflies (Homoptera: Aleyrodidae), are small plant-feeding insects with piercing-sucking mouthparts, and both immature and adult whiteflies feed on the undersides of leaves. Adult whiteflies have the ability to both walk and fly, and females lay eggs either singly in a haphazard manner or in spirals or circles on the undersides of leaves. Whitefly eggs are ovoid and have a peg-like pedicel that is inserted into a slit made by the female’s ovipositor in the leaf surface. Alternatively, eggs may be laid directly into

stomatal openings. A glue-like substance deposited at the base of the pedicel cements eggs in place. The pedicel draws water into the egg from the leaf thereby preventing desiccation before hatching. Most whitefly species are arrhenotokous, and females are produced from fertilized eggs. Males are haploid and eclose from unfertilized eggs. The ratio of male and female whiteflies in a population changes over time and is affected by both temperature and male longevity. Males tend to live for shorter periods and populations appear female biased as a result. Lifecycle The sweetpotato whitefly (Bemisia argentifolii) and greenhouse whitefly (Trialeurodes vaporariorum) may infest vegetable bedding plants. However, greenhouse whitefly is the most common species. Both adult and immature whiteflies have piercing sucking mouthparts, are able to remove fluids and produce honeydew that also results in sooty mold fungus. Winged adult whiteflies are 1/ 16-inch in length, and found on the undersides of the youngest, most tender leaves. Females may lay from 150 to 300 eggs, which hatch into first-instar nymphs in about a week. The crawlers move for a short distance before settling down to feed. After three molts, a pupal stage is formed from which adults emerge in about six days. Whiteflies complete their egg to adult cycle in 21 to 36 days depending upon greenhouse temperatures. Monitoring To monitor whiteflies, check susceptible plants, such as tomatoes, at ten locations in an area of 1,000 square feet, examining plants on each side of the aisle. Look on the undersides of one or two leaves per plant, for nymphs, pupa and adults. Yellow sticky traps can also be used to detect adult whiteflies once populations have reached higher densities. Begin treatments as soon as the first sign of infestation is noted. Fungus Gnats, Shore Flies and Predatory or Beneficial Hunter Flies Lifecycle The damp, moist environment in greenhouses favors both fungus gnats and shore flies. Fungus gnat larvae are translucent, white and legless, about ¼ inch long when mature, and have a shiny black head. The mosquito-like adult

is about 1/8 inch long, with long legs, a pair of clear wings and long antennae. There is a distinct “Y” vein on each wing. Fungus gnats are weak fliers and are frequently observed resting on pot media or running over the foliage or other surfaces. The larvae feed on fungi and decaying organic matter, and often injure seedlings and plants. Larva feeding occurs on young, tender roots and in the stem at the base of the plant. This feeding injury provides an entry for disease pathogens. A female fungus gnat may lay up to 300 whitish eggs in clusters of 20 or more. The eggs are deposited on the surface or in the crevices of moist soil or potting media. Eggs hatch in about six days. Larvae feed for 12 to 14 days before changing into pupae. The pupal stage may last five to six days. Adults live up to ten days. The life cycle from egg to adult requires approximately four weeks depending on greenhouse temperatures. Adult shore flies also occur in damp greenhouses. Shoreflies are often misidentified as fungus gnats or hunters flies but have a distinctly different appearance. The adult shore fly is about 1/8 inch long and has a robust body, very short antennae, shorter legs and dark wings with about five light spots. Adults may be seen resting on plant leaves. Larvae are off-white and do not have distinct head capsules that are characteristic of fungus gnat larvae. Shore flies do not injure plants through direct feeding, but can carry root rot pathogens from diseased to healthy plants. Their fecal spots or droppings can also be unsightly. To manage shore flies, control their food source, algae. Adult hunter flies, a natural enemy (beneficial fly) are also found on sticky cards that may be mistaken for shore flies. Hunter flies can be distinguished from shore flies, by their size and color. Hunter flies are larger, gray in color (males are lighter gray than the females) and do not have light spots on their wings. Hunter flies are in the same family as common houseflies and are similar in appearance, but smaller. Hunter flies may prey upon fungus gnats, shore flies, Liriomyza leafminers, moth flies and whiteflies. Monitoring To monitor for fungus gnat larvae, place raw potato chunks (with peel removed) on the soil surface. Larvae are attracted to the potato chunks and will congregate underneath. Check the potato chunks after 2 days for the larvae. Potato disks cut one inch in diameter and 0.5-1 inch thick are

effective. In addition, choose plants on each bench and inspect the soil surface and around the base of the plant including the stem just below the soil line. Record the location and the level of infestation. Badly infested plants should be removed as they serve as a source of infestation. Adult fungus gnats can be monitored with yellow sticky cards placed at the base of the plant at the soil line. Weekly inspections of yellow sticky cards can detect the onset of an infestation, and continued recording of the number of adults per card per week can aid in evaluating the efficacy of control efforts. Thrips Lifecycle The most injurious species is the western flower thrips (WFT). They often do considerable damage before they are discovered, because thrips are small, multiply rapidly and feed in plant buds in which they can remain undetected. WFT also vector tospoviruses. Feeding marks from the rasping mouthparts of thrips appear as white streaks on the leaves. Infested new growth may curl under and leaves are often deformed. Adult WFT are about 1/16-inch long, with narrow bodies and fringed wings. Females are reddish brown and males are light tan to yellow. The wingless immature larval stages are light yellow. Female thrips insert eggs (several hundred per female) into plant tissue. The tiny yellowish larvae molt twice and feed on plant fluids as they mature. Larvae drop off the plant into the soil and pass through two stages, after which adults emerge. The egg to adult lifecycle can be completed in 7 to 13 days depending upon greenhouse temperature. During warmer temperatures development is more rapid than at cooler temperatures. Monitoring Early detection of a thrips infestation is critical for effective management because populations are lower and it is easier to obtain good spray coverage when plant canopies are small. Symptoms of their feeding are often not noticed until the damage has occurred. Eggplant, tomatoes, and peppers are prone to thrips infestations. Yellow sticky cards, key plants and indicator plants provide an easy way to detect the onset of an infestation. Yellow sticky cards should be placed just above the crop canopy, and near doors, vents and over thrips-sensitive cultivars to monitor the movement of thrips. The light to

medium-blue sticky cards may catch more thrips (and shore flies) than yellow ones. However, it is more practical to use yellow cards for general pest monitoring to attract fungus gnats, whiteflies and winged aphids. The number of thrips per card should be recorded and graphed weekly to monitor population levels and movement in or out of the greenhouse, and thus aid in control decisions. Spider Mites Lifecycle Two-spotted spider mites can be found on vegetable bedding plants. Adult females are approximately 1/50-inch long, and slightly orange in color. All mobile stages are able to pierce plant tissue with their mouthparts and remove plant fluids. Most spider mites are found on the underside of leaves. Feeding injury often gives the top leaf surfaces a mottled or speckled, dull appearance. Leaves then turn yellow and drop. Large populations produce visible webbing that can completely cover the leaves. Eggs are laid singly, up to 100 per female, during her 3 to 4-week life span. Eggs hatch into larvae in as few as 3 days. Following a brief larval stage, several nymphal stages occur before adults appear. Egg to adult cycle can be completed in 7-14 days depending upon temperature. Hot and dry conditions favor spider mite development. Monitoring Check for mites by examining foliage. Adult spider mites are not found on sticky cards. Mites often develop as localized infestations on beans, tomatoes, or eggplants. Sample plants by turning over leaves and with a hands-free magnifier (Optivisor™) or hand lens, check for the presence of spider mites. Cyclamen Mites Life Cycle The shiny, orange-tinted cyclamen mites prefer to hide in buds or deep within the flowers. Adult females can lay from 2 to 3 eggs per day for up to two to three weeks. Eggs are deposited in moist places at the base of the plant. Cyclamen mites can complete their life cycle in 1 to 3 weeks. Females can live up to one month and can reproduce without mating. Cyclamen mite females lay 2 to 3 eggs per day for up to two to three weeks. Cyclamen mite

eggs are oval, smooth and about one half the size of the adult female. Larvae hatch from the eggs in 3 to 7 days. The slow moving white larvae feed for 4 to 7 days. Cyclamen mites prefer high relative humidity and temperatures of 60o F. Cyclamen mites affect a number of ornamental bedding plants including dahlia, fuchsia, gerbera daisy, petunias, viola as well as strawberries in the field. They may migrate to peppers or tomatoes. Monitoring Cyclamen mites pierce tissue with their mouthparts and suck out cell contents. Look for signs of damage which may be concentrated near the buds or occur on the entire plant. Symptoms include inward curling of the leaves, puckering and crinkling. Pitlike depressions may develop. The mite is only 1/100 of an inch long. Examination under a microscope is often needed to confirm the presence of cyclamen mites. Broad Mites Life Cycle Broad mites are closely related to cyclamen mites. They can be distinguished from cyclamen mites by their egg stage. Eggs are covered with “bumps” that look like a row of diamonds. Eggs are best seen using a dissecting microscope. Adults and larvae are smaller than the cyclamen mites and walk rapidly on the underside of leaves. Broad mites can also be attach themselves to whiteflies and use the whiteflies as a carrier for their dispersal. The development of broad mites is favored by high temperatures (70 to 80° F). Broad mites can complete their life cycle in as little as one week. Females lay from 30 to 75 eggs. Monitoring Broad mites can affect a number of ornamentals including gerbera daisy, New Guinea Impatiens, saliva, ivy, verbena and zinnia. They may migrate to peppers or tomatoes. Look for characteristic damage including leaf edges curling downward. Terminal buds may be killed. As they feed, broad mites inject toxic saliva, which results in the characteristic twisted, distorted growth. Broad mite injury can be mistaken for herbicide injury, nutritional (boron) deficiencies or physiological disorders. Inspect the underside of the leaves for the mites and their eggs with a 20x hand lens or submit samples to a laboratory for diagnosis. Microscopic examinations are often helpful.

WEED MANAGEMENT In greenhouses, weeds are a primary source of insects such as aphids, whiteflies, thrips, and other pests such as mites, slugs and diseases. Low growing weeds help maintain moist conditions, a favorable environment for fungus gnats and shore flies. Many common greenhouse weeds such as chickweed, oxalis, bittercress, jewelweed, dandelion and ground ivy can become infected with tospoviruses including impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV) while showing few, if any visible symptoms. Thrips can then vector the virus to susceptible greenhouse crops. Weeds can also carry other plant damaging viruses that are vectored by aphids. An integrated weed management program will help to effectively manage weed populations. This approach includes preventive measures such as sanitation and physical barriers, and control measures such as hand weeding and the selective use of post-emergence herbicides. The use of a physical barrier such as a weed block fabric is an effective method to limit weed establishment on greenhouse floors. The weed fabric should be left bare so it can be easily swept. Covering the weed fabric with gravel makes it difficult to remove any spilled potting media providing an ideal environment for weed growth. Regularly pull any escaped weeds before they go to seed. Repair tears in the weed block fabric. Few herbicides are labeled for use in a greenhouse due to the potential for severe crop injury or death to desirable plants. This injury may occur in a number of ways including: 1) spray drift if fans are operating at the time of application, and 2) volatilization (changing from a liquid to a gas). Herbicide vapors are then easily trapped within an enclosed greenhouse and injure desirable plant foliage. Always be sure the herbicide selected is labeled for use in the greenhouse. Carefully follow all label instructions and precautions. It is the applicator’s responsibility to read and follow all label directions. Use a dedicated sprayer that is clearly labeled for herbicide use only. Avoid use of pre-emergence herbicides in the greenhouse! Preemergence herbicides are applied to soil to prevent the emergence of seedlings. There are currently no pre-emergence herbicides labeled for greenhouse use. They can persist for many months and in some cases over a year. Pre-emergence herbicides can continue to vaporize in the greenhouse, causing significant damage to young transplants. (Note: Surflan (oryzalin) is

no longer registered for use in enclosed greenhouses). Post-emergence herbicides are applied after the weeds have emerged. Several pos-temergence herbicides can be used under greenhouse benches and on the floors. Contact herbicides are best applied to small seedlings. Large weeds will be burned but not killed.

HERBICIDES FOR USE IN GREENHOUSES Clethodim (Envoy Plus): Selective, post-emergence herbicide, for the control of grasses only, works by contact. For use when crops are in the greenhouse. Clove oil (Matratec): Non-selective, post-emergence herbicide. Works by contact. For use when crops are in the greenhouse. Diquat dibromide (Diquat E Pro ZL, Reward): Non-selective. Works by contact. For use when crops are in the greenhouse. Glyphosate (Glyphosate Pro 4, Razor, Roundup Pro, Roundup Pro Concentrate, Touchdown Pro): Non-selective postemergence herbicides. Systemic. For use in an empty greenhouse between crops and outside greenhouses. Pelargonic acid (Scythe): Non-selective, post-emergence, contact herbicide. Cool or cloudy weather may slow down activity. Provides no residual weed control but leaves a strong odor. For use when crops are in the greenhouse. Rosemary oil, clove oil, thyme oil (Sporatec): Non-selective, postemergence herbicide. Works by contact. For use when crops are in the greenhouse. Weed control outside greenhouses In addition to mowing, herbicides may also be used outside of greenhouses. Before spraying weeds around the greenhouse with any herbicide, close windows and vents to prevent spray drift from entering the greenhouse. Avoid using auxin-type herbicides, such as those labeled for broadleaf weed control in turf or brush killers, or herbicides with high volatility near greenhouses. Select herbicides with low volatility.

REFERENCES Cox D. 2016 Plant Response to Nature’s Source and EcoVita Organic Fertilizers vs Plantex Chemical Fertilizer. July-Aug. Floral Notes 29(1). Cox D. 2014. Organic Fertilizers - Thoughts on Using Liquid Organic Fertilizers for Greenhouse Plants. Sept.-Oct. Floral Notes 27(2) Mattson N. Substrates and Fertilizers for Organic Vegetable Transplant Production. Cornell Greenhouse Horticulture, Cornell University. Organic Greenhouse Vegetable Production, Potting Mixes for Certified Organic Production, Organic Greenhouse Tomato Production, Plug and Transplant Production for Organic Systems, ATTRA - National Sustainable Agriculture Information Service. The National Organic Program http://www.ams.usda.gov/nop/NOP/Nophome.html

Guidelines:

16: Plant Parasitic Nematodes Nematodes are some of the most numerous multi-cellular animals on earth. They exist in a vast array of ecologic relationships and may be found in almost every kind of ecologic niche available, from the arctic to deserts or the depths of the oceans. A handful of soil may contain many thousands of these microscopic worms. Most are free-living species that feed on bacteria, fungi, algae or other nematodes. Many are parasites of insects, plants or animals. Most nematodes cannot be seen without a microscope, but some animal parasites may be measured in meters. The nematodes that growers and gardeners are most familiar with and concerned about are the plant parasitic nematodes. They are simple, consisting of only about 1,000 somatic cells in a “tube within a tube” body form. The exterior tube is the outside body wall or cuticle, and the interior tube is the digestive tract that extends from the anterior mouth to the anus near the tail. Plant parasitic nematodes have a stylet, a spear-like mouthpart used to cut into or pierce plant cells. They possess digestive, nervous, excretory and reproductive systems but do not have circulatory or respiratory systems. They cannot see, so they find their way through soil to hosts by means of physical cues and chemical receptors. Plant parasitic nematodes live in water films in soil or in and around plant parts such as roots, stems and leaves. They may be general feeders or have very specific host-parasite relationships with a limited number of host plants. Management of plant parasitic nematodes is challenging in many cropping systems but can be especially challenging in ornamentals for a number of reasons. Herbaceous perennials are a diverse group of about 2,500 species in about 500 different genera. In addition, perennials are propagated by a number of techniques, including seed, division and cuttings. The use of vegetative methods of propagation such as division are often preferred, as they are easier and may produce better, more uniform plants as well as reduce variation within cultivars. Unfortunately, vegetative propagation may result

in the unintended propagation of low populations of plant parasitic nematodes in non-symptomatic plants, increasing the spread and distribution of these nematodes and the extent of nematode problems.

PLANT ROOTS

Fig. 16.1: Root-knot nematode symptoms on carrot

Root-knot nematode symptoms on plant roots are dramatic. As a result of nematode feeding, large galls or “knots” can form throughout the root system of infected plants. Severe infections result in reduced yields on numerous crops and can also affect consumer acceptance of many plants, including vegetables (Fig. 16.1). The degree of root galling generally depends on three factors: nematode population density, Meloidogyne species and “race,” and host plant species and even cultivar. As the density of nematodes increases in a particular field, the number of galls per plant also will increase. Large numbers of nematodes penetrating roots in close proximity also will result in larger galls. Meloidogyne hapla (the northern root-knot nematode) produces galls less than half the size of those produced by M. incognita (the southern root-knot nematode) on the same plant hosts. Finally, each crop responds differently to root-knot nematode infection (Fig. 16.2). Carrots typically undergo severe forking with galling predominantly found on lateral roots (Figures 2, 6). Root-knot nematode galls on lettuce are beadlike. On grasses and onions, galls are usually small and barely noticeable, often no more than slight swellings. Depending upon the crop affected and the severity of

infection, these symptoms can often result in significant economic losses to growers. Most root-knot nematodes have a very widehost range. Thus, growers who have a root-knotnematode problem may find it difficult to controlthe nematode and its damage through croprotation, although this is sometimes a viableoption. Cotton growers who have an infestation of M. incognitacan often plant peanuts in subsequent years to reduce nematode populations. Unfortunately, peanut is an excellent host for a race of M. arenaria race 1, which can be found in fields that also contain M. incognita. Growers who have a problem with M. javanica can employ sweet pepper as a rotational crop, but not if they also have M. incognita. These two examples demonstrate the importance of understanding which Meloidogyne species is present. In addition to differences in pathogenicity on a specific crop, there can be an even greater degree of specialization. For example, M. hapla will not reproduce on grasses. In contrast, M. graminis only reproduces on grasses. Thus growers who experience problems with M. hapla can rotate corn and wheat into traditional vegetable production, provided they have the appropriate equipment available.

Fig. 16.2: Tomatoes grown in soil heavily infested with Meloidogyne.

ABOVE GROUND SYMPTOMS While the most diagnostic root-knot nematode damage occurs below

ground, numerous symptoms can also be observed above ground. Severely affected plants will often wilt readily. Because galled roots have only limited ability to absorb and transport water and nutrients to the rest of the plant, severely infected plants may wilt even in the presence of sufficient soil moisture, especially during the afternoon. Plants also may exhibit nutrient deficiency symptoms because of their reduced ability to absorb and transport nutrients from the soil. Additional fertilization will not generally result in remediation of root-knot nematode-induced chlorosis. Stunting is frequently observed on host crops grown in root-knot nematode-infested fields, and crop yields are reduced. In highly sensitive crops such as lettuce and carrots, initial density of 2 and < 1 eggs/cc soil, respectively, are sufficient to cause economic losses. At high densities, root-knot nematodes can actually kill host plants, particularly if the high populations occur early in the growing season when plants have minimal root mass. Above ground symptoms usually appear on clusters of plants (foci). Because nematodes move slowly through the soil, infestations will gradually radiate outward from an initial point of infection. This can result in large foci of affected plants surrounded by seemingly unaffected plants. Cultivation and other practices that physically move soil and plants will rapidly spread rootknot nematodes.

PATHOGEN BIOLOGY Genus: Meloidogyne Root-knot nematodes were first reported in 1855 by Berkeley, who observed them causing damage on cucumbers. Until Chitwood’s work in 1949, which defined 4 species and one subspecies (M. incognita acrita) within the genus Meloidogyne, the root-knot nematodes were all considered the same species, Heterodera radicola. Researcher described Meloidogyne exigua, the type species of the genus. From this description, Chitwood obtained the name we currently use for the root-knot nematodes. The name Meloidogyne is of Greek origin, meaning “apple-shaped female.” Approximately 100 species of Meloidogyne have been described. The most widespread and economically important species are M. incognita, M. javanica, M. arenaria,M. hapla, M. chitwoodi and M. graminicola. Root-knot nematodes are primarily tropical to sub-tropical organisms, however M.

hapla and M. chitwoodi are well adapted to temperate climates. Like all plant-parasitic nematodes, root-knot nematodes possess a stylet for injecting secretions as well as ingesting nutrients from host plant cells (Fig 16.3, 16.4). Nematodes have no internal skeletal framework, and their “skin” or cuticle acts against internal turgor pressure to maintain body shape and aid locomotion.

Fig. 16.3: A stylet dissected from Meloidogyne hapla

Fig. 16.4: Longitudinal section through the anterior end of a second-stage juvenile of Meloidogyne incognita infecting a red clover root.

Unlike most other plant-parasitic nematodes, root-knot nematode females are globose and sedentary at maturity. They range in length from 400 to 1000 µm. Once they establish a feeding site, they permanently remain at that location within the plant root. The root-knot nematode feeding site is actually a group of cells known as “giant-cells”. When a nematode initially penetrates a plant cell with its stylet, it injects secretory proteins that stimulate changes within the parasitized cells. Parasitized cells rapidly become multinucleate (contain many nuclei) as nuclear division occurs in the absence of cell wall formation. This process is considered to be “uncoupled” from cell division. Cells never actually divide into new cells; they just get bigger and contain more nuclear material. This allows the giant-cell to produce large amounts of proteins which the nematode will then ingest. Giant-cells also act as nutrient sinks, funneling plant nutrients to the feeding nematode. The root-knot nematode does not feed from the cells directly. It forms a feeding tube (from the esophageal gland cell secretions), secreted

from the stylet into the plant cell cytoplasm, which acts as a sieve to filter the cytosol that the nematode ingests. As the name implies, giant-cells can grow very large in size. Triggered by nematode esophageal gland cell secretions, an increase in the production of plant growth regulators has been demonstrated to play a role in this increase in cell size and division. Root cells neighboring the giant-cells also enlarge and divide rapidly, presumably as a result of plant growth regulator diffusion, resulting in gall formation. As the female nematode enlarges, its posterior region may break the epidermis of the root, and the eggs are deposited into a gelatinous egg mass. Mature root-knot females (pearly white in color) can be observed without magnification. Second-stage juveniles (J2) and males can only be observed with the aid of a microscope. Generally, females of root-knot nematodes have a globose body, with a short “neck,” containing their stylet, metacorpus and esophageal gland cells. The J2s of the root-knot nematode are most commonly encountered in soils and are vermiform (worm-shaped). They are usually no larger than 500 µm in length and 15 µm in width. This is the only infective stage. Root-knot nematode males also are vermiform and range from 1100 to 2000 µm in length. They have distinct lips and strongly developed stylets. In addition, they often have visible spicules, for mating, and a blunt, rounded tail. Many Meloidogyne species are parthenogenic or facultatively parthenogenic. This means that males are not necessary to complete the nematode life cycle and viable eggs can be produced by female nematodes in the absence of fertilization. Because of this, males can be rare in a number of species and are only encountered when the nematode population is subjected to an environmental stress.

Fig. 16.5: Perineal pattern excised from a mature Meloidogyne hapla female for identification

Root-knot nematodes can be identified to species using a number of techniques, but one common method is perineal pattern analysis (Fig. 16.5). The perineum (the region surrounding the vulva and anus) of female nematodes displays a pattern of ridges and annulations for each species. While some variation does exist among individuals, these patterns are quite consistent within a species. The analysis of isoenzyme electrophoretic profiles, often using esterase and malate dehydrogenase, is a common method for the diagnosis of Meloidogyne species in properly equipped labs. Likewise, DNA analyses can also be used to identify different species of root-knot nematodes.

DISEASE CYCLE AND EPIDEMIOLOGY

Root-knot nematodes disease cycle

Epidemiology Root-knot nematodes begin their lives as eggs that rapidly develop into J1 (first-stage juvenile) nematodes. The J1 stage resides entirely inside the translucent egg case, where it molts into a J2 nematode. The motile J2 stage is the only stage that can initiate infections. J2s attack growing root tips and enter roots intercellularly, behind the root cap. They move to the area of cell elongation where they initiate a feeding site by injecting esophageal gland

secretions into root cells. These nematode secretions cause dramatic physiological changes in the parasitized cells, transforming them into giantcells. If the nematode dies, so will the giant-cells upon which it feeds.

Fig. 16.6: Vermiform (worm-shaped) male root-knot nematode still contained within the cuticle of the juvenile stage

J2s do not possess reproductive organs. As with all nematodes, root-knot nematodes undergo four juvenile stages, each progressing through a “molting” process similar to that of insects. As a result of this process, juvenile root-knot nematodes have little resemblance to adult males and females. In the J4 stage, the progression from juvenile to globose adult females or to vermiform adult males becomes clearly visible. They emerge as adults from the J4 cuticle (Fig. 16.6). A single female nematode can produce 500 to more than 1000 eggs. The length of a root-knot nematode life cycle varies among species but can be as short as two weeks. Nematodes in cooler regions typically have longer life cycles. Eggs may remain inside root tissue or may be released into the soil matrix. Eggs hatch at random, i.e. hatching does not require exposure to root exudates. Under favorable conditions, root-knot nematode eggs have been reported to survive for at least one year in the soil.

NEMATODE PESTS The northern root-knot nematode, Meloidogyne hapla, is the most important nematode pathogen affecting a wide range of flowering herbaceous perennials. Root-knot nematodes are sedentary endoparasites, meaning that they stay in one place, feeding on nutrient-rich cells in which they initiate inside the root. Juveniles hatch from eggs; infect roots, usually near root tips; establish feeding cells that result in root galls; and develop into swollen females. These females produce a large number of eggs in egg masses that are produced outside the female body. Symptoms of root-knot infection include variously sized galls, excessive root branching and swollen root tips.

Above ground, plants are often stunted, off-color and exhibit symptoms of poor root function. Meloidogyne hapla as this species can readily over-winter and over time, increase in number on perennials in these areas. Plant parasitic foliar nematodes can also cause severe damage to a large number of flowering ornamentals in nurseries and landscape plantings. The range of symptoms observed on flowering ornamentals can vary considerably with plant and nematode species, but leaves, stems, flowers or buds are commonly distorted, dwarfed and killed. There are two different plant parasitic nematodes that can attack above-ground plant parts. The most common foliar nematode, Aphelenchoides fragariae, was listed as able to attack more than 250 plants in 47 families, including many flowering ornamentals. Ditylenchus dipsaci, the stem and bulb nematode, has been found in more than 450 host plants, including both monocots and dicots. Both of these nematodes are widely distributed and can quickly ruin foliage and flowers. Aphelenchoides spp., common name foliar nematodes, are microscopic, wormlike nematodes 0.018-0.047 inches in length. These migratory nematodes usually feed inside foliage. Depending on the host plant, nematode feeding may cause necrotic lesions delimited by veins in crops such as hosta or salvia, or bronzing and discoloration of the foliage in crops such as begonia and anemone. Injured leaves will eventually desiccate, all or in part. Foliar nematodes can survive desiccation for years and may survive without the host or be dispersed in dry, dead tissue. When rehydrated, plant infection occurs after nematode movement in water films or water splash. Nematodes in water films on surfaces that dry quickly are killed; so long periods of leaf moisture increase dispersal and nematode survival during movement. Once inside new leaves, reproduction occurs rapidly. Females lay 25-32 eggs each that can hatch within 3-4 days and mature in 6-12 days, completing the entire life cycle in about two weeks. Ditylenchus dipsaci, the stem and bulb nematodes, are vermiform, and adults are 0.031-0.055 inches long. These nematodes can infect stem, leaf and bud tissues, causing swellings, distortion and necrosis. Stem and bulb nematode infection is often masked by infection as a result of secondary pathogens such as Botrytis, a common secondary fungus on Phlox subulata. These nematodes can live at a wide temperature range, 41-86° F, although optimum temperature is 59-68° F. Ditylenchus females produce 8-10 eggs per

day with a total of 200-500 per female. The juvenile goes through one molt in the egg, hatches and molts three more times before becoming an adult. The entire life cycle can be completed in 19-23 days. Numerous surveys have detected foliar nematodes on annual and perennial flower species worldwide. We have recently recovered the foliar nematodes Aphelenchoides and Ditylenchus from field-grown and potted nursery plants. Aphelenchoides has been a recurring problem on several plant species, and a number of new hosts have been recently identified. Ditylenchus is a particular problem on the flowering perennial Phlox subulata. Both nematodes can increase populations rapidly to tremendous numbers per leaf, and detached or dried leaves can disperse populations. Infection can quickly become widespread and damaging. Both nematodes can survive for long periods of time in dried plant debris. Treatment The concept of initial damage threshold levels, namely that the amount of damage to plant growth is a result of the number of nematodes present at planting, may not apply to nematodes infecting perennials. Low initial nematode densities may greatly increase on susceptible perennial hosts after the planting year and may cause damage over time. Because of this, control of root-knot nematodes in perennials presents a particular challenge. Managing nematodes in high-value ornamentals previously meant using systemic nematicides such as aldicarb or oxamyl. The current lack of labeled nematicide management options requires nursery and landscape nematode management programs based on sanitation and rotation. Sanitation, accomplished by identifying and eliminating M. hapla-infested planting stock and rotating with non-host species, can be effective, especially for field-grown perennials, although the successful use of rotation requires knowledge about the host status of a large number of plant species. The host suitability of M. hapla to nearly 100 common perennials grown in the Northeast has been investigated and published. In addition, we determined that root pruning of bare-root planting stock can greatly reduce or eliminate M. hapla infection and reduce the potential spread of the nematode. Planting resistant plants, such as Rudbeckia fulgida or aster, into M. hapla-infested soils may also reduce or eventually eliminate populations of this nematode. Control of nematodes in foliar plant parts can be extremely difficult.

Diagnosis of plants with foliar nematodes is often confounded by additional infection with fungal pathogens such as Botrytis cinerea. Nursery and greenhouse crops have little or no tolerance, as any nematode-infested material usually results in the establishment and spread of the nematode. Sanitation, avoidance and irrigation water management may help slow the spread of foliar nematodes, but there are no chemical nematicides registered for post-plant nematode control. This lack of control options may be responsible for the recent increased frequency of diagnosed foliar nematode infestations in the Northeast.

BENEFICIAL NEMATODES: FOR BIOLOGICAL CONTROL IN THE GREENHOUSE Nematodes are small, colorless, cylindrical round worms that occur naturally in soils throughout the world. Different species work best against different insect pests. Steinernema feltiae is primarily used against fungus gnat larvae and thrips pupae dwelling in the soil media. Fungus gnat larvae may be parasitized in any larval stage. Nematodes have traditionally been used against soil dwelling pests because they are sensitive to ultra violet light and desiccation. The beneficial nematodes enter the insect host through body openings. These insect killing nematodes multiply within the host and release a symbiotic bacterium (Xenorhabdus spp.) whose toxin kills the target pest, i.e. fungus gnats. The fungus gnat larvae are killed in one to two days by blood poisoning. More than one generation of nematodes may develop in dead host insect in the growing media. The infective juveniles then exit the dead body and search for new hosts to infect. How to use beneficial nematodes The beneficial insect killing nematode S. feltiae is sold under the trade names of NemaShield, Nemasys, Scanmask and Entonem. All of these products are labeled as a soil drench treatment against fungus gnat larvae. Preventative applications to moist soils work best. •

Apply nematodes with a sprayer (remove screens and filters), injector, hose end sprayer or even a watering can in very small operations. • If using an injector, set the dilution to 1:100. Remove all filters or

screens (50 mesh or finer) in any spray lines so that the nematodes can pass through unimpeded and undamaged. • If using a sprayer, keep spray pressure below 300 psi. Although nematodes are applied in water, they are not aquatic animals and therefore they need extra care while in stock and tank solutions. Adequate aeration of the nematode suspension during application is important. This can be done using a small battery powered submersible pump or even mechanically with a stirrer to keep the solution agitated. A small pump will also keep them from settling on the bottom of the stock solution container, which they tend to do. The suspension in the spray tank should be kept cool and applied as soon as possible after mixing. This is especially important during the warmer months. The longer they are kept before spraying and the warmer the tank water, the more quickly their energy reserves are used up. Weaker nematodes are less robust during and after application, and less able to search for and infect a susceptible host. Unlike many traditional pesticides there is no re-entry interval (REI) (an added bonus in propagation houses). No adverse effects have been shown against non-target organisms in many different field studies. But, beneficial nematodes are living organisms, so there are a number of precautions you need to follow for their successful use. Check nematode viability before application • To do this, place a small amount of the product in a small clear container or petri dish. Add 1 or 2 drops of room temperature water; wait a few minutes and look for actively moving or swimming nematodes. They have slight “J” curvature at the end of their bodies. Use a dark black background and a hand lens or field microscope to see the small (0.6 mm or 0.02 inches in length) nematodes. Dead nematodes will straight and still. • Apply in the evening or at dusk or on a cloudy, overcast day. (Nematodes are very sensitive to ultra violet light and desiccation). • Nematodes are compatible with a number of different pesticides. However, they are generally not compatible with organophosphates, carbamates, nematicides and hydrogen dioxide. • Do not mix nematodes with your fertilizer solution!

When you receive the nematodes When you receive the nematodes, check to see that the cold packs are still cold. If you must store the nematodes, store them in a refrigerator at (3842°F). Avoid placing them in a small refrigerator where they may freeze and die! Make sure that there is adequate circulation around each tray or package. Check the expiration date on the package for the length of time they can be stored. Specific Tips when applying Nematodes for Use Against Fungus Gnat Larvae How to tell if the nematodes are working against fungus gnats The symbiotic bacteria break down the host insect’s cuticle. The infected fungus gnat larvae rapidly disappear, so they may be difficult to locate in the growing media. Infected fungus gnat larvae are often opaque-white to light yellow in color. Use potato disks to monitor for fungus gnat larvae. Place disks on the surface of the growing medium two days before application in order to determine the population level prior to treatment, and again 3-5 days and 1012 days after application. Leave the potato disks on the growing media for two days in each case, before examining them for fungus gnat larval activity. Specific tips for use against western flower thrips (from the Nemasys label). • Nematodes require moist conditions to enhance effectiveness. • If plants are dry, provide light overhead irrigation prior to nematode application. • Ensure good foliar coverage of spray mix to enhance contact with the target pest. • Use of a wetting agent or surfactant such as Capsil will enhance wettability of the spray mix and encourage nematode movement. • Following application, ensure that the crop remains wet for at least two hours. Note: Do not apply in direct sunlight. • Note: the nematodes will desiccate after about one day, depending upon environmental conditions. Grower feedback has been variable, with some observing excellent

results and others less so. Efficacy will be variable depending upon the relative humidity, and temperature in your greenhouse, dose applied, frequency of application, and life stage of the thrips. Some growers apply the nematodes with additional water in the summer months to ensure that the foliage stays wet to contact the thrips on the foliage. Depending upon the temperature, relative humidity levels and other environmental conditions, up to 2x the amount of water may be needed to keep the foliage wet for two hours. Regular monitoring, sanitation, proper spacing and judicious use of fungicides and biological fungicides may be needed to discourage foliar diseases. •

Applying the nematodes as a heavy surface spray or “sprench” to young,incoming plant material will have an added benefit of targeting anyincoming fungus gnats in the media as well as thrips pupae. • Growers who have had success with this application method, apply the nematodes on a weekly basis, and target the young growing point where thrips tend to hide. • As with any biological control measure, they are most effectively used preventively in conjunction with good cultural practices for thrips control (sanitation, rigorous weed controls, inspection of incoming plants and regular monitoring). Pest Monitoring and Maximizing Use of Beneficial Insects Many growers no longer rely exclusively on conventional pesticides to control pests. Most have increased their use of mechanical and biological control methods. While pesticide resistance issues have driven much of this change in management philosophy, many growers are now discovering how well biological control agents work and how to easily incorporate them into pest management programs. Beneficial, or insect parasitic, nematodes have become more widely adopted because of their flexibility for tank-mix applications, as they are compatible with other pest control products and insecticides. They are often rotated with insecticides to help manage pests that are or may become resistant. Other beneficial organisms pair well with insect parasitic nematodes,

such as predatory beetles and mites, parasitoids and entomopathogenic fungi to manage insect populations. Development of pest management strategies that marry mechanical, biological and chemical control methods are foundations of integrated pest management (IPM). The growers that see the most success in IPM programs take the time to educate themselves about using beneficial nematodes, and learn how to integrate biological and chemical control methods. The six guiding principles of IPM include : 1. Establishment of action, or economic, thresholds aimed to control, but not to completely eradicate, pest populations. 2.

Regular monitoring, scouting and record keeping to accurately identify and quantify pest populations over time.

3.

Development of cultural practices to maintain healthy plants without excess water, fertilizer and pesticide inputs.

4.

Utilization of mechanical barriers or methods that physically prevent or remove insects from a greenhouse or nursery.

5. Use of biological control agents to manage insect populations. 6. Applying conventional synthetic insecticides responsibly, by only treating when necessary and rotating modes of action to limit potential for targeted pests to develop resistance.

IMPORTANCE OF MONITORING A limitation of many biological control agents is that they cannot quickly control high insect pest populations. Regular monitoring and scouting early in a crop cycle is important to identify the presence of pests. Studying previous pest monitoring and control records will identify what pests were present, at what time of year, what control measure(s) were used, and effectiveness of control measures in controlling the pest. With this knowledge, growers can develop biologically-based IPM programs that minimize the potential of pests reaching economic thresholds. If pests do reach thresholds, conventional insecticides that are compatible with biological control agents can quickly reduce pest populations

to manageable levels. Economic thresholds vary depending on grower preference, crop species and stage, crop susceptibility and growing conditions. Several key practices are recommended for pest monitoring. 1. Schedule time for monitoring and scouting. Scouting and monitoring are essential to a successful IPM program. Scouting should take place once or twice a week throughout the entire production season - no exceptions. 2. Monitor early. Begin monitoring as soon as new plants and cuttings arrive to help identify pests before the populations rise. Quarantine incoming plant material in order to prevent introduction of pests to other areas of the facility. Continue to monitor pest populations throughout the season. 3. Determine infestation levels. Use sticky card counts, potato slices, beating trays and other pest monitoring tools to determine population density and subsequent control measures. • Sticky cards are an inexpensive and effective tool for monitoring flying pests. They help determine when and where the insect appears. • Potato slices attract larval stages of many pests, especially fungus gnat larvae. Place potato slices on the soil surface and check every few days. • Randomly pick up plants while monitoring to look for flightless insects. Study the tops and undersides of leaves and flowers for signs or symptoms of insects. • Some insects will fall from a plant when disturbed. Place a white piece of paper or cloth and tap or gently shake a plant to dislodge insects. Quickly capture, identify and record the dislodged insects. • Look for signs of damage from insect pests. Some insects have chewing mouthparts while others have piercing sucking mouth parts, and insects usually leave characteristic and diagnostic signs. 4.

Keep detailed records. Record the species and number of pests found whilemonitoring. Create a map or outline of areas that had higher infestation levels. Keeping these detailed records helps determine when

certain insects become a problem and when to implement control measures. 5.

Treat preventatively. Because biological control agents work best with low pest populations, it is important to prevent high infestations from occurring. Make regular applications of beneficial nematodes and other pest control methods that you choose. Pest proof netting or other physical barriers often are the first line of defense. When feasible, plants should be selected with resistance to common pests. Likewise, banker or buffer plants also can be effective.

Regular pest monitoring is essential to develop effective IPM programs, and to help guide management and application decisions. Being proactive in controlling insect pests with a combination of biological and chemical control agents will help keep your plants healthy and your customers happy.

DISEASE MANAGEMENT Plant Resistance In certain crops, root-knot nematodes are effectively controlled by resistance genes. In tomato, genetic resistance to root-knot nematodes is conferred by the Mi gene which was obtained from Lycopersicon peruvianum, a wild relative of the common tomato. When resistance genes are transferred into susceptible germplasm, the genetically altered plants become resistant to infection by certain species of root-knot nematode. However, populations of root-knot nematodes that can circumvent root-knot resistance have been identified in both the greenhouse and agricultural fields, suggesting the potential for the eventual failure of root-knot resistance. Many other resistance genes have also been identified that are effective against species of Meloidogyne. These include the Mi2 through Mi8 genes (all from Lycopersicon) and the Me and N genes from pepper. In many cases, however, these genes become ineffective at higher temperatures. Besides these genes, there are a number of genes that have not yet been named, and new sources of genetic resistance to root-knot nematodes are frequently being identified. Biological Control Control measures employing organisms antagonistic to root-knot nematodes have been attempted by many researchers. The most commonly

used biological control agents are fungi and bacteria. There are many kinds of nematophagous (nematode-feeding) fungi. Some fungi use mycelial traps or sticky spores to capture nematodes, for example, Arthrobotrys spp. and Monacrosporium spp. Other fungi parasitize eggs and root-knot nematode females, e.g., Pochonia chlamydosporia and Paecilomyces lilacinus. The major bacterial antagonists are Pasteuria penetrans and species of Bacillus. Endospores of P. penetrans attach to the cuticle of a juvenile nematode, produce penetration structures that enter the nematode, and slowly consume it. Several nematode antagonists have been studied in both greenhouse and field experiments. A number of commercial products based on biocontrol agents are available for the management of root-knot and other nematodes. However, a significant problem in developing effective biological control agents is the inability to economically generate the large amounts of biological material necessary for application over large areas.

INTEGRATED MANAGEMENT The most successful approaches to nematode control rely on integrated pest management strategies (IPM). IPM combines management options to maintain nematode densities below economic threshold levels. IPM techniques can still be difficult to implement against a pathogen as aggressive and resilient as root-knot nematodes. Nevertheless, a combination of management tactics/tools, including cultural practices (rotations with nonhost crops and cover crops that favor the buildup of nematode antagonists), resistant cultivars, and chemical soil treatments, if necessary, generally provide acceptable control of root-knot nematodes. The extent of this success, however, is dependent upon having accurate damage threshold densities and available and readily acceptable resistant cultivars.

REFERENCES Barker, K.R., G.A. Pederson and G.L.Windham. 1998. Plant and Nematode Interactions. ASA, CSSA, SSA Publishers, Madison, WI. France, R.A. and G.S. Abawi. 1993. Interaction between Meloidogyne incognita and Fusarium oxysporum f. sp. phaseoli on selected bean genotypes. J. Nematol. 26: 467-474. Jepson, S.B.1987. Identification of Root-Knot Nematodes (Meloidogyne

Species). CAB International, Wallingford, UK. Karssen,G.2002.The Plant-Parasitic Nematode Genus Meloidogyne Goeldi, 1892 (Tylenchida) in Europe. Brill Academic Publishers, Boston, MA. Lamberti, F. and C.E.Taylor,Eds.1979. Root-Knot Nematodes (Meloidogyne Species). Academic Press, New York. Starr J.L., J. Bridge and R. Cook, eds. Plant Resistance to Parasitic Nematodes. 2002. CABI Publishing, Cambridge, MA. Viaene, N.M.1998. Management of Meloidogyne hapla on lettuce in organic soil with sudangrass as acover crop. Plant Dis. 82: 945-952.

17: Thrips in Greenhouse Crops - Biology, Damage and Management INTRODUCTION Thrips are a major pest of greenhouse crops. A number of thrips species are commonly found including western flower thrips (Frankliniella occidentalis), eastern flower thrips (Frankliniella tritici), onion thrips (Thrips tabaci), and Echinothrips. However, western flower thrips is the predominant species and the most difficult to control. Adult western flower thrips are approximately 1-2 mm in length and generally yellowish-brown in colour. Identification to the species level is difficult (especially among western flower thrips, eastern flower thrips and onion thrips) because they are so small and their colour varies. Adults are the only stage that can be identified to species. Identification should be done by specialists.

Fig. 17.1. Comparison between adult western flower thrips (right) and adult Echinothrips (left).

LIFE HISTORY

The life cycle consists of five stages: egg, larval, prepupal, pupal and adult. Female adult western flower thrips live up to 30 days and lay 2-10 eggs per day. At 20°C, development from egg to adult takes approximately 19 days. At 25°C, it takes 13 days. The eggs are inserted into soft plant tissues, including flowers, leaves, stems and fruit. In sweet pepper, egg hatch gives the leaves a speckled appearance, with the degree of speckling corresponding to the number of hatched eggs. The larval stage (see Fig. 17.2) consists of 2 instars that feed and develop on the leaves, flowers and fruit. The prepupal and pupal stages often complete their development on the ground or growing medium, but pupation can also take place on the plant. The pupa (see Fig. 17.3) is a non-feeding stage during which the wings and other adult structures form. The adults are weak fliers, usually taking short flights from leaf to leaf or plant to plant. Nevertheless, they disperse rapidly throughout the greenhouse. Adult thrips can be transported on wind currents and will enter the greenhouse through vents and doorways. At all stages they may be dispersed on workers’ clothing and on infested plants, growing media or farm implements.

DAMAGE The adult and larval stages feed by piercing the plant surface with their mouthparts and sucking the contents of plant cells. This causes white or brown spots on the leaves where the plant cells have been destroyed. These spots are also speckled with dark fecal droppings from the thrips.

VEGETABLE CROPS

Fig. 17.2. First and second larval instars plus adult of western flower thrips.

Fig. 17.3. Pupal stage of western flower thrips.

In cucumber (see Fig. 17.4) and tomato, thrips damage is noticed first on the lower leaves. In sweet pepper (see Fig. 17.5), it is evident in the upper youngest leaves. Heavy infestations reduce the ability of the plants to photosynthesize, reducing the yield. On vegetable flowers, thrips feeding creates silvery white

Fig. 17.4. Thrips feeding damage on cucumber leaves.

Fig. 17.5. Thrips feeding damage on pepper leaves.

streaks on the petals. Fruit damage varies according to the crop. For instance, in cucumber fruit, feeding creates severe distortion and curling as well as white streaks (see Fig. 17.6). Feeding on sweet pepper (see Fig. 17.7) causes silvery or bronze streaks or spots on the fruit. Thrips also feed on the calyx, causing it to turn up and expose the fruit to bacterial infections. On tomato, thrips may lay eggs in the fruit, creating ghost-spotting (see Fig. 17.8). Ghost-spotting can also occur with sweet pepper and cucumber.

ORNAMENTAL CROPS Western flower thrips has a host range of hundreds of plant species, including many major commercial floriculture crops. Damage includes feeding scars and leaf distortion (see Figs. 17.9 and 17.10). Thrips are particularly attracted to flowers, where they cause damage such as streaking and scarring of petals, distortion of flowers and flower buds and incomplete petal expansion (see Figs. 17.11 and 17.12). Virus Transmission Western flower thrips is the most important vector of a group of viruses called tospoviruses. Tomato spotted wilt virus (TSWV) and impatiens necrotic spot virus

Fig. 17.6. Thrips feeding damage on cucumber fruit.

Fig. 17.7. Egg-laying scars and feeding damage on sweet pepper.

Fig. 17.8. Thrips egg-laying scars on tomato

Fig. 17.9. Thrips feeding damage on roses.

Fig. 17.10. Thrips feeding damage on chrysanthemum leaves.

Fig. 17.11. Thrips feeding damage on chrysanthemum.

(INSV) are the most common tospoviruses in greenhouse crops. The TSWV is generally found in vegetable crops and some ornamental crops such as chrysanthemum, while INSV is more common in ornamental crops. In vegetables, symptoms of this disease vary according to the host, cultivar and stage of plant development, but it can severely reduce or even stop plant growth. Other general symptoms include stunting, bronzing and curling of

the leaves, and distortion of affected plant areas. In addition, infected fruit are misshapen and ripen unevenly, often with a necrotic ring pattern (see Figures 13 and 14). In ornamental crops, many different species serve as hosts for INSV. Symptoms and susceptibility vary widely but include: • • • •

ring spots and line patterns on leaves necrotic lesions black streaking on veins and stems stunting

Fig. 17.12. Thrips feeding damage on gerbera.

Fig. 17.13. TSWV symptoms on pepper fruit.

Fig. 17.14. TSWV symptoms on pepper leaves.

Fig. 17.15. INSV symptoms on kalanchoe: concentric ring patterns.

• death of growing points and crown • plant death in some crops (e.g., gloxinia)

MANAGEMENT Monitoring Monitoring the population levels of western flower thrips is critical for successful pest management. In vegetable crops, monitoring should begin during propagation and continue after transplanting. In floriculture crops, thrips can be present at damaging levels year-round, although populations are usually smaller during winter. Commercially available blue or yellow sticky traps can be used to monitor the population densities of adult thrips (see Fig. 17.16). Blue traps are more attractive to western flower thrips, although yellow traps are more attractive to other pests such as whiteflies and aphids. Your choice depends on how many pests you need to monitor, the susceptibility of the crop to thrips and/or tospoviruses and your need to detect

thrips populations at low levels. When setting up a monitoring program, use 1 trap per 100-200 m2. The exact number will depend on the layout of the greenhouse. A large open range will require a lower total density of cards than a greenhouse made up of a several smaller areas. Place the sticky cards in a grid pattern throughout the greenhouse. Check the traps weekly and record the average number of thrips per trap. Be aware that this is not an absolute measure of the population; rather, it measures increases and decreases in thrips numbers throughout the year. As you become more aware of how the numbers on sticky cards relate to the population in the crop, you can use the monitoring data to help you make pest management decisions. In greenhouse ornamentals, visually inspecting simple flowers, such as impatiens, can provide good estimates of thrips numbers in the crop. However, in more complex flowers, visual counts can be less reliable. In sweet pepper and cucumber crops, precision-level sampling programs have been developed for monitoring adult western flower thrips in the flowers. These sampling programs vary the number of samples taken according to the population level of the pest and accurately predict the pest density to set precision levels.

Fig. 17.16. Sticky cards

Cultural Control Sanitation is the first and most important step in implementing an effective pest management program. Effective sanitation will reduce or even eliminate thrips as a pest problem. For example, in cut roses, removing all flower buds (including non-marketable flowers) can significantly reduce thrips populations in that crop. Cultural control measures also include

maintaining a healthy crop and an optimal greenhouse environment (such as 80% relative humidity), creating less favourable conditions for a rapid increase in the density of thrips populations. Physical Control An influx of outside pests, including thrips, can overwhelm your greenhouse IPM program, making it difficult to plan ahead. To prevent this, use screens to restrict the movement of insects into the greenhouse. Biological control Because thrips have developed resistance to most registered pesticides, biological control is now the primary strategy for controlling thrips in greenhouse crop production. Biological control agents include predatory mites such as: Neoseiulus (= Amblyseius) cucumeris Amblyseius swirskii Iphesius (= Amblyseius) degenerans Stratiolaelaps scimitus (= Hypoaspis miles) Gaeolaelaps gillespiei Gaeolaelaps aculeifer (= Hypoaspis aculeifer) minute pirate bugs (Orius insidiosus) nematodes (Steinernema feltiae) the fungal insect pathogen Beauveria bassiana N. cucumeris and A. swirskii are the most extensively used predatory mites and look very similar. These mites control western flower thrips on the foliage by feeding on the first instar larvae. A. swirskii can also feed to a lesser extent on second instar thrips. As such, it takes a number of weeks for their impact to be seen in the greenhouse, and it is unlikely that they will completely eliminate thrips populations. The life cycle for N. cucumeris is completed in approximately 10 days at 20°C and 6 days at 25°C. A. swirskii has a higher optimal temperature for development than A. cucumeris and performs better in summer conditions. Its development time is similar to that of A. cucumeris but depends on the number and type of prey available. • • • • • • • • •

Predatory mites should be introduced at the beginning of the crop or as soon as thrips are detected. Sanitation at the beginning and end of a cropping

season is extremely important and will delay any thrips infestation until the biological control agents can be effective. Regular introductions of either N. cucumeris or A. swirskii are necessary, either by dispersing bran mixed with mites on plants or growing medium or by hanging a slow-release rearing sachet on plants. The sachet system provides a continuous release of mites to the plant and should be replaced monthly. In ornamental production, many growers are now using new slow-release mini-sachets, which reduce the cost substantially and can be used on individual containers (e.g., hanging baskets or even 15-cm pots). Applying a supplemental food source such as apple pollen to chrysanthemum may help A. swirskii to get established when thrips levels are low. The number of introductions depends on the crop and level of thrips infestation. Control of the thrips should be achieved in 5-9 weeks. When using N. cucumeris or A. swirskii, it is important to maintain at least 70% relative humidity in the greenhouse and avoid using any persistent pesticides for several months before introducing the mites. Orius is effective in controlling thrips. Unlike N. cucumeris and A. swirskii, Orius will feed on all stages of thrips. It is often found in the flowers, where it feeds on pollen as an alternative food source. Because pollen is not often present in ornamental crops, Orius is not as effective in flower crops as it is in vegetables. However, recent research has shown ornamental peppers can be used as a banker plant for Orius in other ornamental crops, allowing a population to establish, develop and disperse within the greenhouse. Some ornamental and vegetable growers are using this strategy to take advantage of the control potential offered by Orius. Development time for Orius from egg to adult is 31 days at 20°C and 19 days at 25°C. Orius enters reproductive diapause when there are less than 12 hr of light per day. Thus, Orius is only effective as a biological control agent from March to September.

Fig. 17.17. Methods for introducing predatory mites: directly on the plants (top), using a bag rearing system (middle) and piling bran on rockwool cubes or other growing medium (bottom).

Orius is best released when the pest level is low. One or two releases are usually enough to provide thrips control in approximately 3-5 weeks, depending on the level of thrips and the type of host crop. For greenhouse vegetable crops, Orius is most successfully used on peppers and cucumber. Introduce adults in several locations where they can naturally disperse by flying throughout the greenhouse. Flower sampling is the best method to monitor the presence of Orius. Iphesius degenerans differs from N. cucumeris and A. swirskii in its appearance and its ability to tolerate less humid conditions. It is dark and very agile. Because it reproduces very well on pollen, it performs best in crops with a pollen source (e.g., greenhouse peppers) but is unlikely to be the best option for floricultural crops. Stratiolaelaps scimitus and Gaeolaelaps gillespiei are soil-dwelling predatory mites that feed on a variety of soil organisms, including thrips pupae. Apply either of these to the growing medium (e.g., rockwool, peat

mixes) once only, at the beginning of the crop. Although it is difficult to determine the exact impact of these predators on a thrips population, research has estimated they can kill up to 30% of pupae. Because they are unlikely to provide enough control on their own, they are better used in combination with other predators. Nematodes are frequently used by ornamental growers in Ontario. Research in Ontario and Europe has shown that they effectively control thrips pupae when applied to the growing medium on a weekly basis. To reduce costs, this is best done by overhead application in propagation, when the plants are pot tight. Beauveria bassiana is a fungal pathogen of thrips. It is usually mixed in water and applied as a spray. Like many fungi, it is more effective under high humidity. Therefore, to treat ornamentals, it is most often applied in propagation. In vegetables, it can be either sprayed onto the crop or distributed via bumble bees that are supplied with hives specially equipped with dispensing trays. These trays containBeauveria bassiana spores that are diluted with a powdered carrier. The bees must walk through the trays to leave the hives. In the process, some of the spore mixture sticks to their bodies. The spores become distributed in the crop when the bees fly in search of nectar and pollen and when they pollinate the crops. When thrips come into contact with spores on the crop surface, they become infected and die. Chemical Control Chemical control of western flower thrips can be difficult. They are resistant to most pesticides and feed deep within the flower head or on developing leaves. This makes them a difficult target for insecticides, so thorough coverage is essential. If you use pesticides to control thrips, follow these general guidelines: •

Begin applications early, before the thrips population grows too large. Thrips are more easily managed when population levels are low. • Although it is important to rotate chemical classes, use only one chemical class for the duration of the thrips’ life cycle. This generally means using a different class every 2-3 weeks, depending on the time of year. • Apply pesticides in early morning or late afternoon, when flight activity of thrips is at a peak. This increases exposure of the thrips to the

pesticides.

18: Monitoring the Greenhouse Environment Growth can be defined as an increase in biomass. The increase in size of a plant or other organism can also be considered as the fundamental definition of growth. The growth of plants is associated with changes in the numbers of plant organs occurring through the initiation of new leaves, stems and fruit, abortion of leaves and fruit, and physiological development of numbers from one age class to the next. Managing growth and development of an entire crop for maximum production involves the manipulation of temperature and humidity to obtain not only the maximum rate of photosynthesis under the given light conditions, but also the optimum balance of vegetative and generative growth of plants for sustained production and high yields. This implies that growers can direct the results of photosynthesis, the production of assimilates, sugars and starches, towards both vegetative and generative in a balance. Generative growth is the growth associated with fruit production. For maximum fruit production to occur, the plant has to be provided both with the appropriate cues to trigger the setting of fruit and the cues to maintain adequate levels of stem and leaf development. The balance is achieved when the assimilates from photosynthesis are directed towards maintaining the production of the new leaves and stems required to support the continued production of fruit. The appropriate cues are provided through the manipulation of the environment, and are subject to change depending on the behavior of the crop. Careful attention must be paid to the signals given by the plant, the indicators of which direction the plant is primarily headed, vegetative or generative, and how corrective action is applied through further manipulation of the environment to maintain high production.

LIGHT

Light limits the photosynthetic productivity of all crops and is the most important variable affecting productivity in the greenhouse. The transpiration rate of any greenhouse crop is the function of three variables; ambient temperature, humidity and light. Of these three, it is light which is usually out of our control as it is received from the sun. Supplementary lighting does offer opportunity to increase yield during low light periods, but is generally considered commercially unprofitable. The other means for manipulating light are limited to screening or shading and are employed when light intensities are too high. However, there are also general strategies to help maximize the crop’s access to the available light in the greenhouse. Properties of Light and its Measurement In order to understand how to control the environment to make the maximum use of the available light in the greenhouse, it is important to know about the properties of light and how light is measured. Considerable confusion has existed regarding the measurement of light (LI-COR Inc.), however it is worthwhile for growers to approach the subject. Light has both wave properties and properties of particles or photons. Depending on how light is considered, the measurement of light can reflect either its wave or particle properties. Different companies provide a number of different types of light sensors for use with computerized environmental control systems. As long as the sensors measure the light available to plants, for practical purposes it is not as important how light is measured, as it is for growers to be able to relate these measurements to how the crop is performing. Light is a form of radiation produced by the sun, electromagnetic radiation. A narrow range of this electromagnetic radiation falls within the range of 400 to 700 nanometers (nm) of wavelength. One nanometer being equal to 0.000000001 meters. The portion of the electromagnetic spectrum which falls between 400 to 700 nm is referred to as the spectrum of visible light, this is essentially the range of the electromagnetic spectrum that can be seen. Plants respond to light in the visible spectrum and use this light to drive photosynthesis. Photosynthetically Active Radiation (PAR) is defined as radiation in the 400 to 700 nm waveband. PAR is the general term which covers both photon terms and energy terms ( LI-COR Inc.). The rate of flow of radiant (light)

energy in the form of an electromagnetic wave is called the radiant flux, and the unit used to measure this is the Watt (W). The units of Watts per square meter (W/m²) are used by some light meters and is an example of an “instantaneous” measurement of PAR (LI-COR Inc.). Other meters commonly seen in greenhouses take “integrated” measurements reporting in units of joules per square centimeter (j/cm²) (LI-COR Inc.). Although the units seem fairly similar, there is no direct conversion between the two. Photosynthetic Photon Flux Density (PPFD) is another term associated with PAR, but refers to the measurement of light in terms of photons or particles. It is also sometimes referred to as Quantum Flux Density (LI-COR Inc.). Photosynthetic Photon Flux Density is defined as the number of photons in the 400-700 nm waveband reaching a unit surface per unit of time (LI-COR Inc.). The units of PPFD are micromoles per second per square meter (micromol/m²). As the scientific community begins to agree on how best to measure light there may be more standardization in light sensors and the units used to describe the light radiation reaching a unit area. Greenhouse growers will still be left with the task of making day-to-day meaning of the light readings with respect to control of the overall environment. Generally speaking, the more intense the light, the higher the rate of photosynthesis and transpiration (increased humidity), as well as solar heat gain in the greenhouse. Of these, it is heat gain which usually calls for modification of the environment as temperatures rise on the high end of the optimum range for photosynthesis, and ventilation and cooling begins. Plants also require more water under increasing light levels. The light use efficiency of plants Plants use the light in the 400 to 700 nm range for photosynthesis, but they make better use of some wavelengths than others. All plants show a peak of light use in the red region, approximately 650 nm and a smaller peak in the blue region at approximately 450 nm. Plants are relatively inefficient at using light and are only able to use about a maximum of 22% of the light absorbed in the 400 to 700 nm region. Light use efficiency by plants depends not only on the photosynthetic efficiency of plants, but also on the efficiency of the interception of light. Maximizing the crop’s access to available light

The high cost of greenhouse production requires growers to maximize the use of light falling on the greenhouse area. Before the crops are able to use the light, it first has to pass through the greenhouse covering, which does not transmit light perfectly. The greenhouse intercepts a percentage of light falling on it allowing a maximum of 80% of the light to reach the crop at around noon, with an overall average of 68% over the day. However, the greenhouse covering also partially diffuses or scatters the light coming into the greenhouse so that it is not all moving in one direction. The implication of this is scattered light tends to reach more leaves in the canopy than directional light which throws more shadows. It is important that the crop be orientated in such a way that the light transmitted through the structure is optimized to allow for efficient distribution to the canopy. Greenhouse vegetable crops have a vertical structure in the greenhouse, so light filters down through “layers” of leaves before a smaller percentage actually reaches the floor. Leaf area index (LAI) is widely used to indicate the ratio of the area of leaves over the area of ground which the leaves cover. Leaf area indexes of up to 8 are common for many mature crop communities, depending on species and planting density. Mature canopies of greenhouse sweet peppers have a relatively high leaf area index of approximately 6.3 when compared to greenhouse cucumbers and tomatoes at 3.4 to 2.3 respectively. The optimum leaf area index varies with the amount of sunlight reaching the crop. Under full sun, the optimum LAI is 7, at 60% of full sun the optimum is 5, at 23% full sunlight, the optimum is only 1.5. This point has application to a growing and developing crop. In Alberta, vegetable crops are seeded in November to December, the low light period of the year. Young crops have lower leaf area indexes which increase as the crop ages. Under this crop cycle, the plants are growing and increasing their LAI as the light conditions improve. Crop productivity increases with LAI up to a certain point because of more efficient light interception, as LAI increases beyond this point no further efficiency increases are realized, and in some cases decreases occur. There is also a suggestion that an efficient crop canopy must allow some penetration of PAR below the uppermost leaves, and the sharing of light by many leaves is a prerequisite of high productivity. Leaves can be divided into two groups; sun leaves that intercept direct radiation and shade leaves, that

receive scattered radiation. The structures of these leaves are distinctly different. The major greenhouse vegetable crops (tomatoes, cucumbers and peppers) are arranged in either single or double rows. This arrangement of the plants and subsequent canopy represents an effective compromise between accessibility to work the crop, and light interception by the crop. For a greenhouse pepper crop, this canopy provides for light interception exceeding 90% under overcast skies and 94% for much of the day under clear skies. There is a dramatic decrease in interception that occurs around noon, and lasts for about an hour when the sun aligns along the axis of north-south aligned crop rows. Interception falls to 50% at the gap centers where the remaining light reaches the ground, and the overall interception of the canopy drops to 80 per cent. The strategies to reduce this light loss would be to align the rows eastwest instead of north-south, reduced light interception occurring when the sun aligns with the rows would take place early and late in the day when the light intensities are already quite low. The use of white plastic ground cover can reflect back light that has penetrated the canopy and can result in an overall increase of 9% over crops without white plastic ground cover. The effect of row orientation varies with time of the day, season, latitude and canopy geometry. It has been demonstrated that at 34° latitude, northsouth orientated rows of tall crops, such as tomatoes, cucumbers and peppers, intercepted more radiation over the growing season than those orientated east-west. This finding was the opposite for crops grown at 51.3° latitude. The majority of greenhouse vegetable crop production in Alberta occurs between 50° (Redcliff) and 53° (Edmonton) North. This would suggest that the optimum row alignment of tall crops for maximum light interception over the entire season, would be east-west. However, in Alberta, high yielding greenhouse vegetable crops are grown in greenhouses with north-south aligned rows as well as in greenhouses with east-west aligned rows. Alberta is known for its sunshine, and the sun is not usually limiting during the summer. In fact, many vegetable growers apply whitewash shading to the greenhouses during the high light period of the year because the light intensity and associated solar heat gain can be too high for optimal crop performance. The strategies for increasing light interception by the

canopy should focus specifically on the times in year when light is limiting, for Alberta, this is early spring and late fall. When light is limiting, a linear function exists between light reduction and decreased growth, with a 1% increase in growth occurring with a 1% increase in light under light levels up to 200 W/m². When light levels are limiting, supplementary artificial lighting will increase plant growth and yield. The use of supplemental lighting has its limits as well. Using supplemental lighting to increase the photoperiod to 16 and 20 hours increased the yield of pepper plants while continuous light decreased yields compared to the 20 hour photoperiod. The economics of artificial light supplementation generally do not warrant the use of supplementary light on a greenhouse vegetable crop in full production. However, supplementary lighting of seedling vegetable plants prior to transplanting into the production greenhouse is recommended for those growers growing their own plants from seed. Light is generally limiting when greenhouse vegetable seedlings are started in November to December. Using supplemental lighting for seedling transplant production when natural light is limiting resulted in increased weight of tomato and pepper transplants grown under supplemental light compared to control transplants grown under natural light. Young plants exposed to supplemental light also were ready for transplanting 1 to 2 weeks earlier than plants grown under natural light. When supplemental lighting was combined with carbon dioxide supplementation at 900 ppm, not only did the weight of the transplants increase, but total yield of the tomato crop was also higher by 10% over the control plants. It is recommended that supplementary lighting be used for production of vegetable transplant production in Alberta during the low light period of the year. This translates to about 4 to 7 weeks of lighting depending on the crop. Greenhouse sweet peppers are transplanted into the production greenhouse at 6 to 7 weeks of age. The amount of light required varies with crop but ranges between approximately 120 - 180 W/m², coming from 400 W lights. A typical arrangement of lights for the seedling/transplant nursery would be to have the lights in rows 1.8 m (6 ft) off the floor, spaced at 2.7 m (9 ft.) along the rows with 3.6 m (12 ft) between the rows of lights . Natural light levels vary throughout theprovince with areas in southern Alberta at 50°latitude receiving 13% more light annually thanareas around

Edmonton at 53° latitude. Strategies tooptimize the use of available light for commercialgreenhouse production involve a number of cropmanagement variables. Row orientation, plantdensity, plant training and pruning, maintaining optimum growing temperatures and relative humidity levels, CO2 supplementation, and even light supplementation, all play a role. All the variables must be optimized for a given light level for a given crop, and none of these variables are independent from one another. How a grower manipulates one variable, affects the others.

Fig. 17.1. High pressure sodium light.

TEMPERATURE MANAGEMENT Development and flowering of plants relates to both root zone and air temperature, and control of temperature is an important tool for the control of crop growth. Managing air temperatures The optimum temperature is determined by the processes involved in the utilization of assimilate products of photosynthesis, i.e. distribution of dry matter to shoots, leaves, roots and fruit. For the control of crop growth, average temperature over one or several days is more important than the day/night temperature differences. This average temperature is also referred to as the 24-hour average temperature or 24-hour mean temperature. Various greenhouse crops show a very close relationship between growth, yield and the 24-hour mean temperature. With the goal of directing growth and maintaining optimum plant balance for sustained high yield production, the 24-hour mean temperature can be manipulated to direct the plant to be more generative in growth, or more vegetative in growth. Optimum photosynthesis occurs between 21 to 22 °C, this temperature serves as the target for managing temperatures during the day when photosynthesis occurs.

Optimum temperatures for vegetative growth for greenhouse peppers is between 21 to 23 °C, with the optimum temperature for yield about 21 °C. Fruit set, however, is determined by the 24-hour mean temperature and the difference in day - night temperatures (Bakker 1989), with the optimum night temperature for flowering and fruit setting at 16 to 18 °C. Target 24-hour mean temperatures for the main greenhouse vegetable crops (cucumbers, tomatoes, peppers) can vary from crop to crop with differences even between cultivars of the same crop. The 24-hour mean temperature optimums for vegetable crops range between 21 to 23 °C, depending on light intensity. The general management strategy for directing the growth of the crop is to raise the 24-hour average temperature to push the plants in a generative direction and to lower the 24hour average temperature to encourage vegetative growth (Portree 1996). Adjustments to the 24-hour mean temperature are made usually within 1 to 1.5 degrees Celsius with careful attention paid to the crop response. One assumption that is made when using air temperature as the guide to directing plant growth is that it represents the actual plant temperature. The role of temperature in the optimization of plant performance and yield is ultimately based on the temperature of the plants. Plant temperatures are usually within a degree of air temperature, however during the high light periods of the year, plant tissues exposed to high light can reach 10 to 12 °C higher than air temperatures. It is important to be aware of this fact and to use strategies such as shading and evaporative cooling to reduce overheating of the plant tissues. Infrared thermometers are useful for determining actual leaf temperature. Precision heat in the canopy Precision heating of specific areas within the crop canopy add another dimension of air temperature control beyond maintaining optimum temperatures of the entire greenhouse air mass. Using heating pipes that can be raised and lowered, heat can be applied close to flowers and developing fruit to provide optimum temperatures for maximum development in spite of the day-night temperature fluctuations required to signal the plant to produce more flowers. The rate of fruit development can be enhanced with little effect on overall plant development and flower set. Precise application of heat in this manner can avoid the problem of low temperatures to the flowers and

fruit which are known to disturb flowering and fruit set. The functioning of pepper flowers are affected below 14 °C , the number of pollen grains per flower are reduced and fruit set under low night temperatures are generally deformed. Problems with low night temperatures can be sporadic in the greenhouse during the cold winter months and can occur even if the environmental control system is apparently meeting and maintaining the set optimum temperature targets. There can be a number of reasons for this, but the primary reasons are 1) lags in response time between the system’s detection of the heating setpoint temperature and when the operation of the system is able to provide the required heat throughout the greenhouse and 2) specific temperature variations in the greenhouse due to drafts and “cold pockets”. Managing root zone temperatures Root zone temperatures are primarily managed to remain in a narrow range to ensure proper root functioning. Target temperatures for the root zone are 18 to 21 °C. Control of the root zone temperature is primarily a concern for Alberta growers in winter, and is obtained through the use of bottom heat systems such as pipe and rail systems. Control is maintained by monitoring the temperature at the roots and maintaining the pipe at a temperature that ensures optimum root zone temperatures. The use of tempered irrigation water is also a strategy employed by some growers. Maintaining warm irrigation water (20 °C is optimum) minimizes the shock to the root system associated with the delivery of cold irrigation water. In cases during the winter months, in the absence of a pipe and rail system, root zone temperatures can drop to 15 °C or lower. The performance of most greenhouse vegetable crops is sub optimal at this low root zone temperature. Using tempered irrigation water alone is not usually successful in raising and maintaining root zone temperatures to optimum levels. The reasons for this are two fold; firstly, the volume of water required for irrigation over the course of the day during the winter months is too small to allow for the adequate sustained warming of the root zone, and secondly, the temperature of the irrigation water would have to be almost hot in order to effect any immediate change in root zone temperature. Root injury can begin to occur at temperatures in excess of 23 °C in direct contact with the roots. The recommendation for irrigation water temperature is not to exceed 24 - 25 °C. The purpose of the irrigation system is to optimize the delivery of water

and nutrients to the root systems of the plants, using it for any other purpose generally compromises the main function of the irrigation system. Systems for controlling root zone temperatures are primarily confined to providing heat during the winter months. During the hot summer months temperatures in the root zone can climb to over 25 °C if the plants are grown in sawdust bags or rockwool slabs, and if the bags are exposed to prolonged direct sunlight. Avoiding high root zone temperatures is accomplished primarily by ensuring an adequate crop canopy to shade the root system. Also, since larger volumes of water are applied to the plants during the summer, ensuring that the irrigation water is relatively cool, approximately 18 °C, (if possible) will help in preventing excessive root zone temperatures. One important point to keep in mind with respect to irrigation water temperatures during the summer months is irrigation pipe exposed to the direct sun can cause the standing water in the pipe to reach very high temperatures, in excess of 35 °C! Irrigation pipe is often black to prevent light penetration into the line which can result in the development of algae and the associated problems with clogged drippers. It is important to monitor irrigation water temperatures at the plant dripline, especially during the first part of the irrigation cycle, to ensure that the temperatures are not too high. All exposed irrigation pipe should be shaded with white plastic or moved out of the direct sunlight if a problem is detected.

MANAGEMENT OF THE RELATIVE HUMIDITY USING VAPOUR PRESSURE DEFICITS Plants exchange energy with the environment primarily through the evaporation of water, through the process of transpiration. Transpiration is the only type of transfer process in the greenhouse that has both a physical and biological basis. This plant process is almost exclusively responsible for the subtropical climate in the greenhouse. Seventy percent of the light energy falling on a greenhouse crop goes towards transpiration, the changing of liquid water to water vapour, and most of the irrigation water applied to the crop is lost through transpiration. Relative humidity (RH) is a measure of the water vapour content of the air. The use of relative humidity to measure the amount of water in the air is based on the fact that the ability of the air to hold water vapour is dependent

on the temperature of the air. Relative humidity is defined as the amount of water vapour in the air compared to the maximum amount of water vapour the air is able to hold at that temperature. The implication of this is that a given reading of relative humidity reflects different amounts of water vapour in the air at different temperatures. For example air at a temperature of 24 °C at a RH of 80% is actually holding more water vapour than air at a temperature of 20 °C at a RH of 80%. The use of relative humidity for control of the water content of the greenhouse air mass has commonly been approached by maintaining the relative humidity below threshold values, one for the day and one for the night. This type of humidity control was directed at preserving low humidity, and although humidity levels high enough to favour disease organisms must be avoided, there are more optimal approaches to control the humidity levels in the greenhouse environment. The sole use of relative humidity as the basis of controlling greenhouse air water content does not allow for optimization of the growing environment, as it does not provide a firm basis for dealing with plant processes such as transpiration in a direct manner. The common purpose of humidity control is to sustain a minimal rate of transpiration. The transpiration rate of a given greenhouse crop is a function of three in-house variables: temperature, humidity and light. Light is the one variable usually outside the control of most greenhouse growers. If the existing natural light levels are accepted, then crop transpiration is primarily determined by the temperature and humidity in the greenhouse. Achievement of the optimum “transpiration setpoint” depends on the management of temperature and humidity within the greenhouse. More specifically, at each level of natural light received into the greenhouse, a transpiration setpoint should allow for the determination of optimal temperature and humidity setpoints. The relationship between transpiration and humidity is awkward to describe, as it is largely related to the reaction of the stomata to the difference in vapour pressure between the leaves and the air. The most certain piece of knowledge about how stomata behave under increasing vapour pressure difference is it is dependent on the plant species in question. However, even with the current uncertainties with understanding the relationships and determining mechanisms involved, the main point to remember about environmental control of transpiration is that it is possible.

The concept of vapour pressure difference or vapour pressure deficit (VPD) can be used to establish setpoints for temperature and relative humidity in combination to optimize transpiration under any given light level. VPD is one of the important environmental factors influencing the growth and development of greenhouse crops, and offers a more accurate characteristic for describing water saturation of the air than relative humidity because VPD is not temperature dependent. Vapour pressure can be thought of as the concentration, or level of saturation of water existing as a gas, in the air. As warm air can hold more water vapour than cool air, so the vapour pressures of water in warm air can reach higher values than in cool air. There is a natural movement from areas of high concentration to areas of low concentration. Just as heat naturally flows from warm areas to cool areas, so does water vapour move from areas of high vapour pressure, or high concentration, to areas of low vapour pressure, or low concentration. This is true for any given air temperature. The vapour pressure deficit is used to describe the difference in water vapour concentration between two areas. The size of the difference also indicates the natural “draw” or force driving the water vapour to move from the area of high concentration to low concentration. The rate of transpiration, or water vapour loss from a leaf into the air around the leaf, can be thought of, and managed using the concept of vapour pressure deficit (VPD). Plants maintained under low VPD had lower transpiration rates while plants under high VPD can experience higher transpiration rates and greater water stress. A key point when considering the concept of VPD as it applies to controlling plant transpiration is the vapour pressure of water vapour is always higher inside the leaf than outside the leaf. Meaning the concentration of water vapour is always greater within the leaf than in the greenhouse environment, with the possible exception of having a very undesirable 100% relative humidity in the greenhouse environment. This means the natural tendency of movement of water vapour is from within the leaf into the greenhouse environment. The rate of movement of water from within the leaf into the greenhouse air, or transpiration, is governed largely by the difference in the vapour pressure of water in the greenhouse air and the vapour pressure within the leaf. The relative humidity of the air within the leaf can be considered to always be 100%, so by optimizing temperature and relative

humidity of the greenhouse air, growers can establish and maintain a certain rate of water loss from the leaf, a certain transpiration rate. The ultimate goal is to establish and maintain the optimum transpiration rate for maximum yield. Crop yield is linked to the relative increase or decrease in transpiration, a simplified relationship relates increase in yield to increase in VPD. Transpiration is a key plant process for cooling the plant, bringing nutrients in from the root system and for the allocation of resources within the plant. Transpiration rate can determine the maximum efficiency by which photosynthesis occurs, how efficiently nutrients are brought into the plant and combined with the products of photosynthesis, and how these resources for growth are distributed throughout the plant. Since the principles of VPD can be used to control the transpiration rate, there is a range of optimum VPDs corresponding to optimum transpiration rates for maximum sustained yield. The measurement of VPD is done in terms of pressure, using units such as millibars (mb) or kilopascals (kPa) or units of concentration, grams per cubic meter (g/m3). The units of measurement can vary from sensor to sensor, or between the various systems used to control VPD. The optimum range of VPD is between 3 to 7 grams/m3, and regardless of how VPD is measured, maintaining VPD in the optimum range can be obtained by meeting specific corresponding relative humidity and temperature targets. Table 18.1 presents the temperature - relative humidity combinations required to maintain the range of optimal VPD in the greenhouse environment. It is important to remember that this table only displays the temperature and humidity targets to obtain the range of optimum VPDs, it does not consider the temperature targets that are optimal for specific crops. There is a range of optimal growing temperatures for each crop that will determine a narrower band of temperature - humidity targets for optimizing VPD. The plants themselves exert tremendous influence on the greenhouse climate , transpiration not only serves to add moisture to the environment, but is also the mechanism by which plants cool themselves and add heat to the environment. Optimization of transpiration rates through management of air temperature and relative humidity can change over the course of the season. Early in the season, when plants are young and the outside temperatures are cold, both heat and humidity (from mist systems) can be applied to maintain temperature and humidity targets. As the season progresses and the crop matures, increasing light intensity increases the transpiration rate and the

moisture content of the air.

Table 18.1. Relative Humidity and Temperature Targets to Obtain Optimal Vapour Pressure Deficits Gram/m3* and millibars (mb) Relative Humidity

‘Optimum range 3-7 grams/m3, 3.9-9.2 mb

To maintain optimum rates of transpiration, venting is employed to reduce the relative humidity in the air. However, under typical summer conditions in Alberta, particularly in the south, ventilation is almost exclusively triggered by high temperature setpoints calling for cooling. Under these conditions, ventilation can occur continuously throughout the daylight period and results in very low relative humidity in the greenhouse. As the hot, moist air is vented, it is replaced by still warm, dry air. Southern Alberta is a dry environment with the relative humidity of the air in summer routinely falling below 30%. Under these conditions some form of additional cooling, mist systems or pad and fan evaporative cooling, is required to both reduce the amount of ventilation for cooling as well as to add moisture to the air. Carbon Dioxide Supplementation Carbon dioxide (CO2) is one of the inputs of photosynthesis and as such CO2 plays an important role in increasing crop productivity. Optimal CO2 concentrations for the greenhouse atmosphere fall with the range of between 700 to 900 ppm (parts per million) (Romero-Aranda et al 1995, Tremblay and

Gosselin 1998). Crop productivity depends not only on efficiency of interception of light but also on the efficiency with which light is converted to chemical energy in photosynthesis. Carbon dioxide enrichment to 1200 ppm increases the maximum conversion efficiency by a substantial amount (between 28 to 59%) (Wilson et al 1992). Photosynthetic efficiencies appear never to exceed about 22 % of the absorbed light energy in the 400 to 700 nm range, the maximum efficiency is obtained at relative low light intensities, not in brightest sunlight. Considering the supply of light to available land area on which a crop is growing, the overall yield efficiencies are always much below 22%. The use of CO2 in greenhouses can give light use efficiencies exceeding those of field crops (Wilson et al 1992). Glasshouse crops with CO2 enrichment achieve maximum efficiency of light energy utilization between 12-13%. The ability of plants to utilize CO2 is dependent upon the presence of light, for this reason it is only useful to supplement CO2 during the daylight hours. The key enzyme for CO2 fixation is rubisco. The activity of rubisco depends on the ratio of the O2 and CO2 concentration in the atmosphere. The major effect of CO2 enrichment is the shift in balance in the O2 and CO2 ratio which improves the activity of rubisco. The effect is just as important at low as at high light levels since the percentage effect on relative growth rate is about the same over a range of light levels. Transpiration rates are reduced under CO2 enrichment conditions by 34%. Increased net leaf photosynthesis rate and decreased transpiration rate under CO2 enrichment is well documented. One of the most important effects of CO2 enrichment is the increased water use efficiency. The technique of enriching the greenhouse atmosphere with CO2 to maximize yield is standard practice. The largest increase in growth rate achieved with CO2 enrichment is obtained with high light intensities. A high CO2 concentration may partially compensate for low light levels. There is obviously a potential for synergism between CO2 and light, however the relationship between CO2and light conditions may be relatively loose. When greenhouse ventilation rates are high, the cost of CO2 supplementation can rise steeply. This is particularly so with a ventilation

regime where ventilation is triggered at temperatures between 19-21 °C. Investigations into delaying ventilation to increase the cost effectiveness of CO2 supplementation have shown that the amounts of CO2 supplied to the greenhouse could be reduced by 23 to 35% while still maintaining the CO2 content of the greenhouse atmosphere above ambient CO2 concentrations. Delaying ventilation to conserve CO2 resulted in higher greenhouse temperatures with fruit temperatures exceeding 30 °C. However, total marketable yield fell by 11% and the proportion of fruit graded as Class 1 was reduced by 20% on average. The best advice for CO2 supplementation under high ventilation rates is to maintain the CO2 concentration at or just above the normal ambient level of approximately 350 ppm. This is a highly efficient way of using CO2 supplementation. Maintaining the CO2 concentration at the same level as ambient, there can be no net exchange of CO2 with the outside air through leakage or ventilation. For practical purposes, the input of CO2 is therefore equal to that being assimilated by the crop during photosynthesis, i.e. the utilization of supplementary CO2 is totally efficient. The main point being that ventilation and economical CO2 enrichment may be applied simultaneously. At higher temperatures, 25 °C, net photosynthesis begins to decline and the supplementation of CO2 above this temperature is not considered cost effective (Portree 1996). During longer periods of elevated CO2 the stomata remain partially closed and the reduction of transpiration may cause insufficient cooling, hence, heat damage to the leaves under conditions of intense light. However, the increased VPD associated with the higher temperatures has been shown to counteract the effect of stomatal closure due to CO2 supplementation. Since young plants grow nearly exponentially, they can benefit more from optimal growing conditions than mature plants. Carbon dioxide enrichment results in heavier transplants and can be used to accelerate the growth, as well as improving the quality of the transplants. Carbon dioxide may increase sugar translocation in the roots as well as facilitating the movement of nitrogen and carbon compounds directed towards the

development of new roots. In short, CO2 supplementation shortens the nursery period and results in sturdier, higher quality plants.

AIR POLLUTION IN THE GREENHOUSE Air pollutants can be a concern for greenhouse production. The incidence of air pollutant injury to plants is increasing as more growers use double plastic greenhouses, or other forms of greenhouse sealing to reduce energy loss. Air pollutants can cause visible injury to the leaves, can reduce growth rates or both. Tomatoes and cucumbers are particularly sensitive to air pollutant injury. When considering the effects of greenhouse air pollutants ,it is important to remember that these pollutants pose significant health risks for people working the crops. Common pollutants are often by-products of combustion. Although sources of pollutants can be outside the greenhouse, a number of sources of pollutants can be found within the greenhouse. Pollutants can be produced by direct-fired heating units, gas supply lines or carbon dioxide generators that burn hydrocarbon fuels such as natural gas. Significant sources of pollutants outside the greenhouse can include industrial plants or vehicle exhaust.

Table 18.2. Maximum acceptable concentration (ppm) of some noxious gases for humans and plants Gas

Humans

Plants

Carbon Dixoide (CO2)

5,000

4,500

Carbon monoxide (CO)

47

100

Sulfur dioxide (SO2)

3.5

0.1

Hydrogen sulfide (H2S)

10.5

0.01

Ethylene (C2H4)

5.0

0.01

Nitrous oxide (NO)

5.0

0.01 to 0.1

Nitrogen dioxide (NO2)

5.0

0.2 to 2.0

Air pollution from sources within the greenhouse commonly arise through cracked heat exchangers on furnaces or incomplete combustion in

the furnace or CO2generators. Heaters and generators should be checked at the beginning of the cropping season to ensure they are operating properly and complete combustion is occurring. The most common air pollutants resulting from incomplete combustion include nitrogen oxides, nitric oxide (NO) and nitrogen dioxide (NO2), sulfur dioxide (SO2), ethylene (C2H4), propylene (C3H6), ozone (O3), carbon monoxide (CO) and hydrogen sulfide (H2S). Symptoms of air pollutant injury vary with the specific gases involved. The common symptoms of sulfur dioxide injury is characterized by severe leaf burn appearing within 24 to 36 hours of exposure to high levels of the gas. There is a distinct line between the affected and unaffected areas on the leaves and young leaves are more susceptible to injury than mature leaves. Symptoms of NO2 injury include darker than normal green leaves with downward curling leaf margins and dead areas on the leaves in severe cases. Ethylene functions as a plant growth regulator, involved in seed germination, root development, flower development and leaf abscission. Ethylene injury can include a reduction in growth, shortening and thickening of stems and twisting of stems, as well as premature leaf and flower drop. Propylene injury is similar to ethylene but usually occurs at concentrations 100 times higher than those for ethylene. Ozone injury is characterized by mottling, necrotic flecking or bronzing necrosis of leaves, premature leaf drop and decreased growth. Growing Media Most commercial vegetable production greenhouses in Alberta use some form of “hydroponic culture”. The term hydroponics essentially translates as ‘water culture’. It is an advanced form of crop culture which allows for specific control of the delivery of nutrients to the plants. The term hydroponics can bring to mind a number of variations on the same theme. Hydroponic growing systems can include: substrate culture where the roots are allowed to grow in an inert or semi-inert media; solution culture where the roots are immersed in ponds of nutrient solution; NFT culture (nutrient film technique) where the roots are contained such that a thin film of nutrient solution constantly runs by the roots; and aeroponics where the root systems are suspended within an enclosed area and are misted with nutrient solution. A general working definition of hydroponic culture that would include all of

the above systems, is plant culture where the plants receive fertilizer nutrients every time they receive water. Using this working definition of hydroponics also leaves room for the inclusion of soil as a growing medium. However, soil culture is not widely practiced in commercial vegetable greenhouses in Alberta. The main reason for moving out of soil, into soilless culture, is to escape problems due to soil borne diseases that can build-up in the soil used year after year. Soilless media such as rockwool and sawdust offer an initially disease-free growing medium. There are other advantages of moving the root system out of the soil and into confined spaces such as sawdust bags or rockwool slabs. The main advantages are realized in the improved management of watering and nutrition, topics which are discussed in more detail in following sections. Media for seeding and propagation Rockwool plugs are the most common media used for seeding. Rockwool is manufactured by subjecting rock mineral materials to very high temperatures and then spinning the materials into a fibre. The plugs can be square (2 cm × 2 cm by 4 cm deep) and can come joined together as a rockwool “flat” that fit into standard 28 cm × 54 cm plastic seeding flats. As the seed germinates and the seedlings are ready for their first transplanting, the plugs easily separate from each other when the seedlings are transplanted into rockwool blocks. Rock wool blocks are typically around 10 cm × 10 cm by 8 cm deep, with a depression cut into the upper surface to receive the rockwool plug at the first transplanting. As the seedling continues to grow, the root system develops from the rockwool plug into the confines of the block. When the seedling is ready for transplanting into the main production greenhouse at “house set”, the bottom of the rockwool block is placed in direct contact with the larger volume of growing media used in the production house. Growing media for the production greenhouse The majority of commercial greenhouse vegetable production is based on substrate culture where the plants are grown in sawdust or rockwool. These substrates contain practically nothing in the way of plant nutrients and serve as a substrate for the root system to anchor the plant. The growing media plays a significant role in defining the environment of the root system and

allows for the transfer of water and nutrients to the plant. Typically, for sawdust culture, 2 or 3 plants are grown in 20 to 25 litre white plastic bags (white reflects more light) filled with spruce and/or pine sawdust. Rockwool culture uses approximately 16 litres of rockwool substrate for every 2 to 3 plants. The sawdust bags or rockwool slabs are placed directly on the white plastic floor of the greenhouse. Sawdust is less expensive than rockwool in initial cost, however standard density rockwool slabs can be pasteurized and reused for up to three years. Sawdust is a waste product of the lumber milling process which is usually burned, so the use of sawdust as a growing media is an environmentally sound practice. For sawdust culture it is important to use a moderately fine sawdust, lumber mills understand the sawdust requirements for plant production and will supply “horticultural grade” sawdust if they are made aware that the sawdust is to be used for plant culture. Using sawdust that is too fine will break down over the production season with resulting loss of airspace around the roots which can lead to root death. There is always some decomposition of the sawdust during the growing season which makes the product useful for further composting or adding to mineral soils to improve soil quality. Through the continued action of soil microbes the sawdust residue at the end of the cropping season is returned to the environment in an ecologically sound manner. The waste from sawdust culture is confined to the plastic bags themselves which are recovered when the sawdust bags are dumped and can be recycled where facilities exist.

MANAGEMENT OF IRRIGATION AND FERTILIZER FEED In hydroponic crop production systems the application of water is integrated with the application of the fertilizer feed. The management of fertilizer application to the plants is therefore integrated with the management of watering. The management of watering and nutrition is focused on the optimal delivery of water and nutrients over the various growth stages of the plant, through the changing growing environment over the production year, in order to maximize yield. Water quality

Plants are comprised of 80 to 90% water and the availability of adequate quality water is very important to successful crop production. The quality of water is determined by what is contained in the water at the source; well, dugout, town or city water supply, and the acidity or alkalinity of the water. Water is a solvent, and as such, it can contain or hold a certain quantity of soluble salts in solution. Fertilizers, by their nature, are soluble salts, and growers dissolve fertilizers in water to obtain nutrient solutions in order to provide the plants with adequate nutrition. Prior to using any source of water for crop production it is important to have it tested for quality. Water quality tests determine the amount of various salts commonly associated with water quality concerns. The maximum desirable concentrations, in parts per million (ppm), for specific salt ions in water for greenhouse crop production are presented in Table 18.3. Parts per million are one unit of measurement of the amount of dissolved ions, or salt in water, and are also used to measure the level of dissolved fertilizer salts in nutrient solutions. The level of nutrients as dissolved ions in water can also be reported in milligrams/Litre of solution. There is a direct relationship between milligrams/Litre (mg/L) and ppm, where 1 mg/L = 1 ppm. Another common unit of measure for dissolved fertilizer salts is the millimole (mM), the concept of millimoles and the relationship between millimoles and ppm is explained in the special topic section.

Table 18.3. The maximum desirable concentrations, in parts per million (ppm), for specific salt ions in water for greenhouse crop production Element

Maximum desirable (ppm)

Nitrogen (NO3 - N)

5

Phosphorus (H2PO4 - P)

5

Potassium (K+)

5

Calcium (Ca++)

120

Magnesium (Mg++)

25

Chloride (Cl-)

100

Sulphate (SO —) 4

200

Bicarbonate (HCO -) 3

60

Sodium (Na++)

30

Iron (Fe+++)

5

Boron (B)

0.5

Zinc (Zn++)

0.5

Manganese (Mn++)

1.0

Copper (Cu++)

0.2

Molybdenum (Mo)

0.02

Fluoride (F-)

1

pH

75

E.C.

1

1 mmho/cm = 1 mS/cm = 1000 microsiemens/cm Water quality tests will also report the pH, the acidity or alkalinity of the water. Once the source of water has been determined as suitable for greenhouse crop production it is also important to have the water tested routinely to ensure that any fluctuations in quality that may occur does not compromise crop production. Electrical conductivity of water Water quality analyses also report the electrical conductivity or E.C. of the water. The ability of water to conduct an electrical current is dependent of the amount of ions or salts dissolved in the water. The greater the amount of dissolved salts in the water, the more readily the water will conduct electricity. Electrical conductivity is an indirect measurement of the level of salts in the water and can be a useful tool for both determining the general suitability of water for crop production, and for the ongoing monitoring of the fertilizer feed solution. Using electrical conductivity as a measure to maintain E.C. targets in the nutrient solution and the root zone can be used as a management tool for making decisions regarding the delivery of fertilizer solution to the plants. Electrical conductivity is measured and reported using a number of measurement units including millimhos per centimeter (mmhos/cm), millisiemens per centimeter (mS/cm) or microsiemens per centimeter. Water suitable for greenhouse crop production should not have a E.C. in excess of 1.0 mmhos/cm.

PH The relative acidity and alkalinity of the water is expressed as pH, and is measured on a scale from 0 to 14. The lower the number, the more acidic the water or solution, the higher the number the more alkaline. The pH scale is a logarithmic scale, meaning that every increase of one number ie. 4 to 5, represents a ten times increase in alkalinity. Conversely, every single number decrease, ie. 5 to 4, represents a ten times increase in acidity. Most water supplies in Alberta are alkaline, with typical pH levels of 7.0 to 7.5. Alkalinity of the water increases with increasing levels of bicarbonate. The pH measurement reflects the chemistry of the water and nutrient solution. The pH of a fertilizer solution has a dramatic determining effect on the solubility of nutrients, how available the nutrients are to the plant. The optimum pH of a feed solution, with respect to the availability of nutrients to plants, falls within the range of 5.5 to 6.0. The pH of a solution can be adjusted through the use of acids such as phosphoric or nitric acid, or potassium bicarbonate, depending on which direction the feed solution needs to be adjusted. When acids or bases are used to adjust the pH of the feed solution, the nutrients added by the acid; nitrogen, phosphorus, must be accounted for when the feed solution is calculated. Most water supplies in Alberta are basic in pH and require the use of acid for pH correction. The amount of acid required to adjust the pH is usually dependent on the bicarbonate (HCO3-) level in the water. The amount of bicarbonate in the water supply can be determined by a water analysis, and is reported in ppms. A good target pH for nutrient feed solution is 5.8, and as a general rule this pH corresponds to a bicarbonate level of about 60 ppm. If the incoming water has, for example, a pH of 8.1 and a bicarbonate level reported at 207 ppm, 207 ppm - 60 ppm = 147 ppm that needs to be neutralized by acid to reduce the pH from 8.1 to 5.8. In order to neutralize 61 ppm, or 1 milliequivalent, of bicarbonate it takes about 70 ml of 85% phosphoric acid, or about 84 ml of 67% nitric acid per 1000 litres of water. In order to neutralize 147 ppm of bicarbonate: Using 85% phosphoric acid 140 / 61 = 2.3 milliequivalents of bicarbonate to be neutralized

2.3 milliequivalents x 70 ml of 85% phosphoric acid for each milliequivalent = 2.3 x 70 ml = 161 mls of 85% phosphoric acid for every 1000 litres of water. Using 67% nitric acid 2.3 milliequivalents of bicarbonate to be neutralized. 2.3 milliequivalents x 76 ml per milliequivalent = 2.3 × 76 ml = 175 mls of 67% nitric acid for every 1000 litres of water These calculations have to be made for each water sample based on the results of water a analysis reporting the level of bicarbonates. In addition to phosphoric and nitric acid, sulfuric and hydrochloric acids can also be used to adjust the pH of the water down. Acids are corrosive. Special care and attention must be used when handling them for pH correction. The common acids used to lower the pH are phosphoric acid (85%) and nitric acid (67%), of these two, nitric acid is the most corrosive and must be handled very carefully. Acid resistant safety glasses, rubber gloves and a rubber apron should be the minimum safety equipment used when handling acids.

THE MINERAL NUTRITION OF PLANTS In order to support optimum growth, development and yield of the crop, the fertilizer feed solution has to continually meet the nutritional requirements of the plants. Although the mineral nutrition of plants is complex, experience in crop culture has determined basic requirements for the successful hydroponic culture of plants. There are 13 mineral elements that are considered essential for plant growth. Water (H2O) and carbon dioxide (CO2) are also necessary for plant growth and supply hydrogen, carbon and oxygen to the plants bringing the total to 16 essential elements. A criterion to determine whether an element is essential to plants is if the plant cannot complete its life cycle in the complete absence of the element. In addition to the essential elements there are other elements, although not necessarily considered universally essential, which can affect the growth of

plants. Sodium (Na), chloride (Cl) and silicon (Si) are in this category, all three of these nutrients either enhance the growth of plants, or are considered essential nutrients for some plant species. The essential nutrients can be grouped into two categories reflecting the quantities of the nutrients required by plants. Macronutrients or major elements, are required by plants in larger quantities, when compared to the amounts of micronutrients, or trace elements required for growth. Another useful grouping of the mineral nutrients is based on the relative ability of the plant to translocate the nutrients from older leaves to younger leaves. Mobile nutrients are those which can readily be moved by the plant from older leaves to younger leaves, nitrogen is an example of a mobile nutrient. Calcium is an example of an immobile nutrient, one which the plant is not able to move after it has initially been translocated to a specific location.

Table 18.4. The essential mineral elements for plants Element

Symbol

Type

Mobility in Plant

Symptoms of Deficiency

Nitrogen

N

macronutrient

mobile

Plant light green, lower (older) leaves yellow.

Phosphorus

P

macronutrient

mobile

Plant dark green turning to purple.

Potassium

K

macronutrient

mobile

Yellowish green margins on older leaves.

Magnesium

Mg

macronutrient

mobile

Chlorosis between the veins on older leaves first, turning to necrotic spots, flecked appearance at first.

Calcium

Ca

macronutrient

immobile

Young leaves of terminal bud dying back at tips and margins. Blossom end rot of fruit (tomato and pepper).

Sulfur

S

macronutrient

immobile

Leaves light green in color.

Iron

Fe

micronutrient

immobile

Yellowing between veins on young leaves (interveinal chlorosis), netted pattern.

Manganese

Mn

micronutrient

immobile

interveinal pattern

Boron

B

micronutrient

immobile

Leaves of terminal bud becoming light green at bases, eventually

chlorosis,

netted

dying. Plants “brittle.” Copper

Cu

micronutrient

immobile

Young leaves dropping, wilted appearance.

Zinc

Zn

micronutrient

immobile

interveinal chlorosis of older leaves.

Molybdenum

Mo

micronutrient

immobile

Lower leaves pale, developing a scorched appearance.

The discussion of plant nutrients as elements does not allow for a more complete discussion of how plants access the elements from the root environment, and how hydroponic growers ensure that their crop plants are adequately supplied with nutrients. The term “element” can be defined as a substance that cannot be broken down into simpler substances by chemical means, the basic unit of an element is the atom (Boikess and Edelson 1981). With the simplest, or purest form of plant nutrients being the atom, nutrients are not often available to plants in their purest form. Pure nitrogen is an example of a nutrient element represented by its atom. When the atoms of different elements combine, they can form other substances which are based on a particular combination of atoms, substances based on molecules. Nitrate (NO3-), is a molecule based on nitrogen and oxygen atoms, nitrate is absorbed by plant roots as a source of nitrogen. Nitrate is an “available” form of nitrogen. The nitrate molecule has an overall negative charge, which causes the molecule to be fairly reactive chemically, and therefore more available. The availability of nutrient elements to plants is generally based on the existence of the nutrient element as a charged particle, either a charged atom or charged molecule. An atom or molecule that carries an electric charge is called an ion, and positively charged ions are called cations, while negatively charged ions are called anions. The nitrate molecule (NO3-) is an anion, the iron atom can exist as the Fe+2 (ferrous) or Fe+3 (ferric) cations (Boikess and Edelson 1981). Plants are able to acquire the essential mineral elements via the root system utilizing the chemical properties of ions, particularly that to acquire negatively charged anions, the plant roots have sites that are positively charged. The plant is also able to attract positively charged cations to negatively charged sites on the root. Water is a very important component in the acquisition of nutrient elements by the plants as the nutrient ions only exist when they are in

solution, when they are dissolved in water. As solids, the ions generally exist as salts, a salt can be defined as any compound of anions and cations. In the absence of water, the nutrient ions form compounds with ions of the opposite charge. Anions combine with cations to form a stable solid compound. For example, the nitrate anion (NO3-) commonly combines with the calcium (Ca+2) or potassium (K+) cations forming the larger calcium nitrate Ca(NO3)2 potassium nitrate (KNO3) salt molecules. As salts are added to water, they dissolve, or dissociate into their respective anion and cation components. Once in solution they become available to plants. An important point to remember is that different salts have different solubilities, that is, some salts readily dissolve in water (highly soluble), and some salts do not. Calcium sulfate (CaSO4) is a relatively insoluble salt and would be a poor choice as a fertilizer because very little of the calcium would go into solution as the calcium cation (Ca++) and be available to plants. Fertilizer salts, by their very nature, are useful because they go into solution readily. In hydroponic culture, greenhouse growers formulate and make a water based nutrient solution by dissolving fertilizer salts. In addition to existing as salts, some of the micronutrients; iron, zinc, manganese and copper, exist in chelates. A chelate is a soluble product formed when certain atoms combine with certain organic molecules. The sulphate salts of iron, zinc, manganese and copper are relatively insoluble and chelates function to make these mineral nutrients more readily available in quantity to the plants.

FERTILIZER FEED PROGRAMS Fertilizer nutrient solutions are formulated to meet the needs of the plants using a combination of component fertilizer salts. The amounts of the various fertilizers used are dependent on target levels which have been determined to be optimal for the crop in question. Although there is considerable similarity between fertilizer programs for the various vegetable crops, there can be some differences reflecting the different requirements of the crop. In any event, when mixing fertilizer solutions, only high quality water-soluble fertilizers should be used.

Table 18.5. Forms of mineral nutrient elements that are commonly available to plants Element

Symbol

Available as

Symbol

Nitrogen

N

Nitrate ionAmmonium ion

NO -NH + 3 4

Phosphorus

P

Monovalent phosphate ion

H PO 2 4

Divalent phosphate ion

HPO -2 4

Macronutrients

Potassium

K

Potassium

K+

Calcium

Ca

Calcium ion

Ca+2

Magnesium

Mg

Magnesium ion

Mg+2

Sulfur

S

Divalent sulfate ion

SO -2 4

Chlorine

Cl

Chloride ion

Cl-

Iron

Fe

Ferrous ionFerric ion

Fe-2Fe-3

Manganese

Mn

Manganous ion

Mn+2

Boron

B

Boric acid

H3BO4

Copper

Cu

Cupric ion chelateCuprous ion chelate

Cu+2Cu+

Zinc

Zn

Zinc ion

Zn+2

Molybdenum

Mo

Molybdate ion

MoO 4

Micronutrients

The required nutrient levels, or target nutrient levels of the various essential elements are often expressed as the desired parts per million (ppm) in the final nutrient solution. The recommended nutrient fertilizer feed targets for greenhouse peppers are listed in Table 18.6. Even though all thirteen mineral elements are essential for plant growth and development, nutrient targets for sulfur and chlorine are not listed. The reason for this is adequate amounts of sulfur are obtained from the use of sulfate fertilizers, potassium sulfate or magnesium sulfate. Chloride is assumed to be present in adequate amounts as a contaminant in a number of fertilizers. As the purity of fertilizers has improved, growers will have to pay more attention to ensuring these other elements, particularly chloride, are present in adequate amounts. Once the recommended nutrient targets are known, calculations are made to

determine how much of each fertilizer is required in order to meet them. In order to make these calculations some other basic information is required: 1. The volume of water that will be used to make the feed solution. 2. The types of fertilizers that are available, and the relative amounts of each nutrient present in the fertilizer.

Table 18.6. Nutrient feed targets (ppm) for greenhouse sweet peppers grown in sawdust Nutrient

Target (ppm)

Nitrogen

200

Phosphorus

55

Potassium

318

Calcium

200

Magnesium

55

Iron

3.00

Manganese

0.50

Copper

0.12

Molybdenum

0.12

Zinc

0.20

Boron

0.90

Every greenhouse must be able to supply water and nutrients on an ongoing basis. During hot, dry Alberta summers, mature pepper plants can use approximately 3.5 to 4.0 litres of water per plant per day, cucumbers can require over 6 litres, tomatoes up to 3 litres. This water always contains fertilizer which is added as the water comes into the greenhouse and before it is pumped to the plants. There are a number of variations on the theme, but some form of fertilizer injection system is used in all commercial scale greenhouses. Feed targets and plant balance The first approach to altering the feed solution in response to a crop that is overly vegetative is to increase the feed E.C. to direct the plants to become more generative and set and fill more fruit. The feed EC can be increased

from 2.5 mmhos to approximately 3.0 mmhos over the course of a few days. Dialing up the feed E.C. increases the absolute amounts of fertilizer nutrients in the feed but does not affect the ratio of the nutrient levels with respect to one another. Increasing the feed E.C. increases the level of fertilizer salts in the root zone, increasing the stress on the plant as it becomes more difficult for the plant to take up water. The plant responds to the stress by putting more emphasis on fruit production, a stressed plant begins preparations for the end, by trying to ensure that the next generation will carry forward. The fruit holds the seed, and in plant terms, developing fruit means that the next generation will survive and carry on. Now plants don’t think these things out, but stressing the plant does direct the plant to set more fruit. There is a limit to how far growers can go with this as a successful crop requires having enough vegetative growth to continually fill a high volume of fruit consistently throughout the season. The N:K ratios presented in the table, are all about 1:1.5, increasing the level of potassium, with respect to nitrogen, and increasing the ration to 1:1.7 will direct the plant to be more generative. The reason for this is that nitrogen promotes vegetative growth while potassium promotes mature growth, generative growth. Calcium is also important for promoting strong tissues, fruit, and mature growth. Shifting the feed program to favor potassium over nitrogen will direct the plant to be generative. Calcium is important in the equation in that it should always be approximately equal to the amount of nitrogen. A N:Ca ration of 1:1, works for both tomatoes and peppers, while a N:Ca ratio of 1:0.85 has shown to work well for cucumbers. Changes to the N:K ratio should be made carefully, the above ratios come from the feed programs of successful Alberta growers and can serve as a guide. The place to start is to determine the ratios in the current feed program and examine the performance of he crop. If it is determined that there is room for improving the balance of the plants, alterations in the nutrient ratios can be undertaken. Always be aware that many factors influence plant balance: day/night temperature split, 24 hour average temperature, relative humidity and watering. These factors should be optimized before feed ratios are changed. You have to know where the crop is in order to make sound decisions on where it should be, and how to get there.

Due to the large volumes of fertilizer feed solution that can be required daily, it is impractical to make the fertilizer feed on a day to day basis. Instead, the required fertilizers can be mixed in a concentrated form, usually 100 to 200 times the strength that is delivered to the plants. Injectors or ratio feeders are then used to “meter-out” the correct amount of fertilizer into the water which make up the nutrient solution going to the plants. By using concentrated volumes of the fertilizer feed held in “stock tanks” growers are able to reduce the number of fertilizer batches they have to make. Depending on the number of plants in the crop, the size of the stock tanks, and the strength of the concentrate, growers may only have to mix fertilizer once every 2 to 4 weeks.

DESIGNING A FERTILIZER FEED PROGRAM The design of a fertilizer feed program is a relatively straight forward process once the nutrient target levels are decided and basic information about the water quality, feed delivery system, and component fertilizers are known. Fertilizer targets and the component fertilizers used to make the fertilizer solution can change over the course of the year depending on the crop and the knowledge of the grower. Often the changes are slight adjustments in the relative proportion of the macronutrients to one another, particularly the nitrogen:phosphorus:potassium (N:P:K) ratio. Changes can also include the addition of alternate forms of a nutrient in question, a common example is the use of ammonium nitrogen (NH4 - N) in addition to nitrate nitrogen (NO3 - N) during the summer months. Ammonium nitrate is the common source of ammonium nitrogen, which is a more readily available form of nitrogen that works to promote vegetative growth. During the summer months a target of approximately 17 ppm of ammonium nitrogen is recommended to help optimize plant balance and crop production. Moles and millimoles in the greenhouse; Just another couple of rodents? - Just when you thought you had all your rodent problems under control, some greenhouse vegetable growers have been concerned about millimoles and moles. Not to worry, these growers are not referring to four legged moles. Rather they are using another unit of measure to discuss fertilizer feed targets and root zone targets. So, what exactly is a millimole? A millimole is one thousandth of a

mole, and a mole is defined as the amount of a substance of a system which contains as many elementary entities as there are atoms in exactly 12 grams of 12 C (Carbon 12). Now, you were probably expecting that a definition would help clarify the situation, isn’t that what definitions are supposed to do? The concept of the mole has come out of stoichiometry, that branch of chemistry which studies the quantities of reactants and products in chemical reactions. Now a lot of chemists and physicists have argued for a long time over how to measure the masses of individual elements (some of those same elements that growers feed their crops in fertilizer feed solutions) and in 1961 they settled on using the mole. A good way to understand what a mole is and why to use it is to related it to the concept of a dozen. We understand that a dozen is twelve of something, be it cucumbers, eggs or whatever. A mole is 6.02 x 1023 of some entity, and chemists usually refer to actual molecules of a substance when they talk about moles, although you could have a mole of eggs or a mole of cucumbers. You would be quite the grower to grow a mole of cucumbers, tomatoes or peppers. The number 6.02 × 1023, which in long hand is 602,000,000,000,000,000,000,000, is called Avogadro’s number after the nineteenth century chemist who did some pioneering work on gases and was largely ignored for his trouble. The lesson here is that if you do something great and are not feeling appreciated for the greatness, someone, far into the future may name a big number after you. Moles do relate to parts per million (ppm), they are both ways to measure how much of a given nutrient we are dealing with in a fertilizer feed sample, leachate or tissue sample. The difference is that ppm is a measure of mass (e.g. 1 ppm = 1 milligram/litre) and moles measure amounts. One mole of any substance contains Avogadro’s number of entities or basic units. Those entities, as mentioned earlier, can be atoms or molecules or whatever you want. When we talk about one mole of nitrate nitrogen, NO3, we are referring to 6.02 × 1023 molecules of NO3, because the basic NO3 entity is made up of one atom of nitrogen (N) and three atoms of oxygen (O). If we are talking about a mole of iron, Fe, we are talking about atoms, because the basic entity of iron is the iron atom. All atoms and molecules have different basic weights, some being heavier than other. If we talk about 1 ppm of NO3 versus 1 ppm of Fe, we are

talking about the same mass of each, i.e., 1 milligram/litre. However, there will be a different number of basic entities or moles of NO3 and Fe in a solution which contains 1 ppm each of NO3 and Fe. Now, we are getting close to being able to convert ppm to moles or millimoles, but we will first consider the concept of atomic and molecular weights. The atomic weights of all the elements can be found on the periodic table, that handy chart that we carried with us throughout all our chemistry classes. The atomic weights of the elements are given in grams per mole. The molecular weight of oxygen is 16 grams/ mole, this means that 6.02 × 1023 atoms of oxygen weights 16 grams. One mole of nitrogen weighs 14 grams. By combining all the atoms which make up molecules we can arrive at the molecular weights. Therefore, the molecular weight of NO3 would equal 14 + 3(16) grams/mole or 62 grams/mole. One last thing to remember is that moles are related to millimoles the same way that grams are related to milligrams. So if moles are related in the range of grams, millimoles are in the range of milligrams. We know that 1 ppm is equal to 1 milligram/litre, so to convert ppm to millimoles you divide ppm by the molecular weight of the element you are working with. For example: • 1 ppm of NO3 = 1 mg/litre 1 mg/ litre of NO3 / 62 mg/mole = 0.016 millimoles of NO3 in one litre • 1 ppm of Fe = 1 mg/litre 1 mg/litre of Fe / 56 mg/millimole = 0.018 millimoles of Fe in one litre. • 1 ppm of magnesium (Mg) = 1 mg/litre 1 mg/litre of Mg / 24 mg/millimole = 0.041 millimoles of Mg in one litre. As these examples show, a solution containing 1 ppm of various elements or molecules will contain different mole or millimole amounts of these same elements. To convert millimoles to ppm: ppm = millimoles/litre x molecular weight (mg/millimole) Example:

ppm NO3 = 0.016 millimoles of NO3 in one litre x 62 mg/millimole = 1 ppm NO3 Once you can work back and forth between ppms and millimoles, you might be asking if there is any benefit to working in millimoles rather than ppm. If you are comfortable working with ppms and you are comfortable with designing and managing your fertilizer feed programs in ppms, stick to what you know. However, if you want to be working with actual amounts of atoms and molecules of the nutrients you are feeding then you may want to work with millimoles. Whatever the case, with a little practice you can work with either unit. Calculating the required amounts of the various fertilizers is dependent on the volume of water to be used. This is determined by the volume of the stock tank (e.g. 200 litres) multiplied by the injection ratio (e.g. 100:1 or 200:1). For example, using a 200 litre fertilizer concentrate stock tank, and a 200:1 injection ratio, the volume of water that will be used to calculate the amount of fertilizer to add will be: 200 litres (stock tank volume) x 200 (injector ratio) = 40,000 litres Accounting for the nutrients present in the raw water Assuming the water quality analysis has determined that the water is suitable for greenhouse crop production, the first step is to account for the nutrients that are already contained in the water. This information comes directly from the water analysis report. Accounting for the nutrients provided by the pH adjustment of the water Next determine if the pH needs adjusting, and if so, the amount of acid (or base) required to meet the target pH of 5.8. Once the amount of acid to be added has been determined, the levels of nutrients present in the acid have to be accounted for. Using the example in the previous section, where it was determined that 161 ml of 85% phosphoric acid would be required to adjust the pH from 8.1 to 5.8 for every 1,000 litres of water, the amount of acid required for 40,000 litres would be (161 ml/ 1,000 litres x 40,000 litres =) 6,440 mls. Knowing the volume of acid required, and the specific gravity of the acid, it is possible to calculate the weight of acid that will be used.

Table 18.7. The specific gravity of 85% phosphoric and 67% nitric acid. Phosphoric acid (85%)

1.41 grams/ml

Nitric acid (67%)

1.28 grams/ml

6,440 mls (85% phosphoric acid) × 1.41 grams/ml = 9,080 grams of phosphoric acid. Having the weight of the acid, it is now possible to determine the amount of phosphorus contributed to the pH-adjusted water by 85% phosphoric acid. One more piece of information is required, phosphoric acid contains 32 % available phosphorus. This is also referred to as the fertilizer grade of the acid. Now, using the following formula:

the amount of phosphorus (in ppm) contributed by 7,857 grams of 85% phosphoric acid

This same sequence of calculations can be used to determine the amount of nitrogen contributed if 67% nitric acid was used. In this example 49 ppm of nitrogen would be contributed if 67% nitric acid was used. Determining the required amounts of the various fertilizers necessary to meet the feed targets For the purposes of this discussion of designing a fertilizer program, only component fertilizers will be considered, a list of the common component fertilizers for greenhouse crop production is presented in Table 18.9. The fertilizers are identified by their chemical name, and their fertilizer number designation, ie. 0-53-35 for monopotassium phosphate. The “grade” of the fertilizer with respect to the different nutrients supplied by the fertilizer, is also provided. It is important to know that the three number designation of the fertilizer represents the percentages or grade of nitrogen (N), phosphorus (P) and potassium (K), in that order, that is present in the fertilizer. However, it is very important to note when the percentages for phosphorus and

potassium are used, the number on the bag represents the percentages of phosphate (P2O5) and potash (K2O) and not actual phosphorus and potassium. Phosphate is only 43% actual phosphorus and potash is only 83% actual potassium. For this reason, monopotassium phosphate, 0-53-35, is listed as containing 23% phosphorus (53% x 0.43 = 23%), and 29% potassium (35% x. 0.83 = 29%).

Table 8. Fertilizer “upgrades” of phosphoric and nitric acid Phosphoric acid

32% available phosphorus(PO4-P)

Nitric acid

22% available nitrogen(NO3-N)

Blended, or premixed fertilizers are also used by some growers. A common premixed fertilizer is 20-20-20. If these fertilizers are used it is important to account for all the nutrients provided in the fertilizer, both macro and micronutrients. As well, although the fertilizer 20-20-20 contains 20% nitrogen, for the purposes of calculating actual phosphorus (P) and actual potassium (K), 20-20-20 should actually be considered as 20-8.6-16.6. In determining the amount of fertilizer to add, it is important to remember that as salts, fertilizers often contain more than just one plant nutrient. For example, calcium nitrate (Ca(NO3)2) provides both calcium and nitrogen. Calcium nitrate is commonly used in commercial vegetable greenhouses as the only source of calcium. The amount of calcium nitrate added depends on how much is required to meet the calcium target. However since nitrogen is also present in calcium nitrate, it is important to keep track of how much nitrogen is contributed. After all, there is also an optimum target for nitrogen. Calcium nitrate is 19 % calcium and 15.5 % nitrogen so for every 100 grams of calcium nitrate there will be 19 grams of calcium and 15.5 grams of nitrogen. The percentage of the relative nutrient components of a fertilizer is also sometimes referred to as the “grade.” As the fertilizer calculations are made, an ongoing tally is kept on what nutrients are being supplied by the various fertilizers until all the feed targets have been met. With the information of stock tank size, injector ratio, and the nutrients contributed by each fertilizer, the same relatively simple formula an be used to determine the amount of each fertilizer required to meet the parts per million (ppm) feed targets of the essential nutrients.

This formula can be rearranged to calculate ppm if the amount of fertilizer added is known.

Table 18.9. Some component fertilizers for formulating nutrient feed programs for hydroponic greenhouse vegetable crops Fertilizer Nutrients Macron utrients Nitrogen

Calcium nitratel5.5-0-0

15.5% nitrogen (N03-N)19% calcium

Potassium nitratel 3-0-44

13% nitrogen (N03-N)37% potassium

Ammonium nitrate34-0-0

17% nitrogen (N03-N)17% (NH4-N)

Phosphorus

Monopotassium 53-44

23% phosphorus29% potassium

Potassium

Potassium nitratel 3-0-44

37%potassium13% nitrogen (N03-N)

Potassium sulfateO-0-50

41.5% potassium17% sulfur

Monopotassium 53-44

23% phosphorus29% potassium

Calcium

Magnesium

Sulfur

Chlorine

phosphateO-

phosphateO-

Potassium chlorideO-0-60

49% potassium26% chlorine

Calcium nitratel5.5-0-0

19% calcium15.5% (N03-N)

Calcium chlorideCaCI2-2H20

27% calcium48% chlorine

Magnesium 7H20

10% magnesium13% sulfur

sulfateMgS04-

Magnesium nitrateMg(N03)26H2

10% magnesium11% nitrogen (N03-N)

Magnesium 7H20

10% magnesium13% sulfur

sulfateMgS04-

Potassium sulfateO-0-50

41.5% potassium17% sulfur

Calcium chlorideCaCI2-2H20

27% calcium48% chlorine

Potassium chlorideO-0-60

49% potassium26% chlorine

Iron chelate

13% iron

Micronutrients Iron

nitrogen

Manganese

Manganese chelate

13% manganese

Copper

Copper chelate

14% copper

Molybdenum

Sodium molybdate

39% molybdenum

Boron

Borax

15% boron

Continuing with the example, using 200 Litre stock tanks, a 200:1 injector ratio, meeting a calcium target in the nutrient solution of 180 ppm, and obtaining all the calcium from calcium nitrate. The formula can be used to determine the amount of calcium nitrate required to meet the calcium target, as well as determining the levels of nitrogen (in ppm) contributed by the calcium nitrate.

RULES FOR MIXING FERTILIZERS Once the amounts of the various fertilizers have been determined, the next step is to mix the fertilizers in the stock tanks. Most commercial vegetable greenhouses use a two stock tank system for mixing fertilizers, although some systems involve three stock tanks with the third tank containing the acid or bicarbonate for pH adjustment. Before mixing fertilizers ensure that a dust mask and gloves are worn to avoid inhaling the fertilizer dusts or contacting the fertilizer concentrates. The first rule in mixing fertilizers is to always use high quality, water soluble “greenhouse grade” fertilizers. Second, when working with stock tank concentrates, never mix calcium containing fertilizers (e.g. calcium nitrate) with any fertilizers containing phosphates (e.g. monopotassium phosphate) or sulfates (e.g. potassium sulfate, magnesium sulfate). When fertilizers containing calcium, phosphates or sulfates are mixed together as concentrates the result is insoluble precipitates of calcium phosphates and calcium sulfates. Essentially the calcium combines with the phosphate or sulfate in the solution and comes out of the solution as a solid. This solid forms a “sludge” at the bottom of the fertilizer tank which can plug the irrigation lines. This reaction between calcium, phosphate and sulfate can be avoid if a 1-times strength fertilizer is being mixed, as it is considerably more dilute. However, mixing fertilizers to make a 1-times strength fertilizer solution is

impractical for a commercial greenhouse operation as it would necessitate that someone be mixing fertilizers almost continuously. The third rule for mixing fertilizers is to dissolve the fertilizers for each tank together in hot water. The components of tank 1 are dissolved together as are the components of tank 2. The micronutrients are added to the tanks when the solution is warm, not hot. Fourth, is to continually agitate the solution in the stock tanks as the fertilizers are being added. Using the two-tank stock tank system, the fertilizers should be mixed as follows: Tank A

Tank B

calcium nitrate potassium nitrate (one half the total amount)

potassium nitrate (one half the total amount) magnesium sulfate

iron chelate

monopotassium phosphate potassium sulfate manganese chelate zinc chelate copper chelate sodium molybdate boric acid

If other fertilizers are used, ensure that mixing calcium containing fertilizers with phosphate or sulfate containing fertilizers is avoided. Generally other nitrate fertilizers can be added to the “A” tank, while with all others mixed in the “B” tank. Note that iron is always added to the “A” tank to avoid mixing it with phosphate fertilizers, which can cause the precipitation of iron phosphates (Wieler and Sailus 1996), resulting in iron deficiency in the plants. If acids are used for pH correction, they are generally added to either the “A” or “B” tank or can be added to a third stock tank a “C” tank. If potassium bicarbonate is required for pH correction, it should be added to a third tank, the “C” tank to avoid the risk of raising the pH in the other stock tanks which could result in the other fertilizers coming out of solution. The fertilizer feed program is designed to supply specific quantities of the nutrient elements to the plants per every unit volume of nutrient feed delivered to the plant. The absolute quantities of these nutrients is measured by the parts per million (ppm) targets. In addition to the absolute quantities of

the nutrients in the feed, the relative ratios of one nutrient to another (particularly the N:P:K ratio) is also an important component of the feed program. Direct measurement of the various component nutrients contained in the feed solution, and the determination of the relative ratio of the nutrients comes from a lab analysis of the feed solution. It is useful to have the feed solution tested regularly in order to monitor the actual nutrient levels being delivered to the plants. Lab analysis of the feed solution takes time and it is also important to be able to monitor the feed on a ongoing basis throughout the day. Measuring the electrical conductivity (E.C.) of the feed solution is a very useful tool in the day-to-day management of the fertilizer feed solution. Measurement of the E.C. of the fertilizer feed solution delivered to the plants can be used as an indirect measure of the level of nutrients reaching the plants. The feed program contains the appropriate quantities of dissolved fertilizer salts required to meet the nutrient requirements of the plants, and this solution has a corresponding E.C. In fact, the corresponding E.C. of most feed solutions delivered to the plants, when based on a nitrogen target of 200 ppm, is about 2.5 mmhos. Of course the other nutrients are present in their relative amounts with respect to nitrogen. Once the feed solution has been mixed to meet the targets, measuring the E.C. of the 1-times strength solution can serve as the point of reference for delivering the nutrients to the plants. The day-to-day management of the delivery of feed to the crop can vary and is based on the salt level of the feed solution. The feed solution can be used as a management tool to direct the development of the crop towards a vegetative or generative direction. The basis for this is the higher the level of salts delivered to the root zone, the more stress that is placed on the plants. The more stress that the plant is under, the more emphasis the plant puts on producing fruit and the less emphasis on stems and leaves. There are limits to the salt stresses that can be placed on the plants while still maintaining optimum production, as a high sustained yield is obtained through a balance of leaves and fruit throughout the season. However, using the feed solution to help optimize plant balance is a management tool. On cloudy days, plants can make use of higher fertilizer levels, than on sunny days where the plant has greater demands for water. Raising the feed E.C. on a cloudy day will provide more nutrients to the plants, lowering the fertilizer E.C. on a sunny day will provide a greater relative proportion of water to the plants. The saltier the fertilizer solution, the harder the plant has to work to extract the

water from the root zone. Management of the daily application of fertilizer to the crop is based on varying the E.C. of the feed solution. The general rules for managing the feed E.C. and the total amount of nutrient solution volume delivered to the crop on a daily basis is presented in the next section. Application of Fertilizer and Water Water and fertilizer are delivered simultaneously to the crop via the nutrient solution, and the amounts of water and fertilizer delivered varies with the changing requirements of the plants. The plant’s requirements change as they develop from seedlings to mature plants and in accordance with the day to day changes in the growing environment. In order to manage the delivery of nutrients and water to the plant, it is important to have a way of determining the crop’s requirements for fertilizer and water. Feed monitoring stations are established throughout the crop, one or two stations per every 0.4 hectare (1 acre) of greenhouse area are usually sufficient, but having one monitoring station for every watering “zone” of the greenhouse is a good idea. The purpose of the monitoring station is to measure the volume of feed delivered to the individual plants, and to determine the volume of feed solution leachate, or “over-drain” that is flowing past the plants and out of the root zone over the course of the day. The E.C. and pH of the feed solution is taken on a daily basis, as is the E.C. and pH of the leachate. Daily monitoring the percentage of feed solution volume flowing through the root zone environment, the sawdust bags, or rockwool slabs etc., is used to adjust the volume of feed solution delivered to the plants. The E.C. of the leachate is used to make adjustments on the feed solution E.C. Monitoring the pH of the feed and leachate helps to ensure that the correct pH is being fed to the crop and gives an indication of what is happening in the root zone with respect to pH. Optimum feed pH is approximately 5.8, and this pH optimum also applies to the root environment as well. Generally the activity of the roots tends to raise the pH in the root environment and feeding at a lower pH can help counteract this rise in pH. It is not recommended to feed at a pH of lower than 5.5 when attempting to lower the pH in the root zone. In addition to feeding at a pH of 5.5, the use

of ammonium nitrate at 2 to 5 ppm of ammonium nitrogen (NH4 - N) will help to lower the pH of the root zone due to the acidifying effect of this fertilizer. A schematic of a typical feed monitoring station is presented in figure 19. In addition to monitoring the feed and leachate, recording the leachate percentage, feed and leachate E.C. and pH can be used as a tool to chart the performance of the crop with respect to these recorded values over time, and in relationship to other parameters including the amount and intensity of available light. The amount of nutrient solution delivered to the plant on a daily basis can be determined by the percentage over-drain or leachate that is recovered from the plants over the course of the day. Leaching, or allowing a certain percentage of nutrient solution applied to the crop, to pass through the root system, allows for a flushing of the root zone to avoid the accumulation of salts. Generally, when the plants are young, a percentage leachate of 5 to 10% is a good target. As the plants develop, the amount of water required to attain this over-drain target increases. As the season progresses and the light levels increase and the plants mature and begin to bear fruit, the over-drain targets increase to 20 to 30%. Generally these higher over-drain targets apply as the high light period of the year begins, usually in June. As the percentage over-drain decreases, the leachate E.C. increases, that is, the amount of salts in the root zone increases. The general rule for managing the level of salts in the root zone is that the root zone E.C. should not be greater than 1.0 mmho above the feed E.C. The design of the feed solution is based on delivering adequate nutrition to the plants, and these feed programs usually have an E.C. 2.5 mmhos (this is largely dependent on the E.C. of the irrigation water). With the optimum feed solution E.C. at approximately 2.5 - 3.0 mmhos, the salt levels in the root zone should be maintained at around 3.5 - 4.0 mmhos. Early in the crop cycle, the salt levels in the root zone can be maintained at the proper target fairly easily by increasing the volume of nutrient solution delivered to the plant to ensure a 5 to 10% over-drain. As the season progresses and the water has been increased so that the upper limit of 30% over-drain has been reached, and the E.C. of the over-drain continues to climb above the target of 3.5 - 4.0 mmhos, the E.C. of the solution can be dropped. The reduction in feed solution E.C. is accomplished in stages with gradual, incremental

reductions in feed E.C. in the order of 0.2 mmhos every 2 to 3 days. It is never advised to apply straight water to the plants in order to lower the root zone E.C., since the rapid reduction in root zone E.C. and increased pH can reduce the performance of the crop and compromise the health of the roots (Maree 1994). During periods when the plants are in a rapid stage of growth, the E.C. in the root zone can be below that of the feed solution. For example, the feed can be at 2.5 mmhos while the leachate E.C. may be a 2.0 mmhos. This is an indicator that the plants require more nutrients and the feed E.C. should be increased in increments in the order of 0.2 mmhos until the E.C. in the root zone begins to approach the upper target limit of 4.0 mmhos. By varying the volume and E.C. of nutrient solution delivered to the plants, in accordance to the leachate over-drain and E.C. targets, it is possible to optimize the delivery of adequate water and nutrients to the crop without over watering and over fertilizing. Applying too much or too little water can compromise the health and performance of the crop. The delivery of water to the plants occurs over the course of the entire day. Watering can be scheduled by using a time clock or in more sophisticated systems the watering events can be triggered by the amount of incoming light received by the greenhouse. In general, the greater the ability to control the delivery of water, the greater the ability to maximize crop performance. A starting point for watering the crop early in the crop cycle would be to apply water every half hour from one half hour after sunrise to approximately one hour before sunset. The amount of water required to meet the over-drain target is divided amongst the waterings based on the duration of the individual waterings. For example if a 40 second watering delivers 100 ml of water, then 10 watering events are required to deliver one litre of water. When more than a litre of water is required in one day the duration of the individual watering events can be increased, or the number of watering events can be increased or both. Generally, as the crop matures, it is better to increase the frequency of watering events than the duration of each event. If the watering system allows the variation of the frequency and duration of the watering events over the course of the day, then it is possible to increase the frequency and/or duration of the watering events during the high light period

of the day without necessarily increasing the duration of the early morning or late afternoon watering events. Watering frequency can be used to help direct the vegetative/generative balance of the plant. For any given volume of water that is delivered to the plants, the more frequent the waterings throughout the day, the more the plant will be directed to grow vegetatively. The longer the duration between waterings, the stronger the generative signal sent to the plant. Frequent watering during the summer months in Alberta can help balance plants that are overly generative due to the intense sunlight, high temperatures and low relative humidity. When the concept of percent over-drain is discussed, it is preferable to obtain the majority of the over-drain during the high light period of the day. The first of the over-drain should start to occur at 10:00 am and the greater part of the daily over-drain target should be reached by 2:00 to 3:00 p.m. Having the capability of varying the duration of the watering events over the course of the day allows for more nutrient feed being delivered to the plants between 10:00 am and 2:00 - 3:00 p.m. The use of over-drain targets is one way to ensure the plants are receiving adequate water throughout the day. Another strong indicator of whether or not the plants have received adequate water during the previous day is whether the growing points, or the tops of the plants have a light green color early in the morning. Over the course of the day when the plant is under transpiration stress, the color of the plants will progress from a light green to a darker blue-green. If the plants have received adequate water throughout the previous day, the light green color will return overnight as the plant recovers and improves its water status. If the plants remain a darker bluish-green in the early morning, the amount of water delivered the previous day was inadequate. Usually, this means that the over-drain target for the previous day have not been met and the amount of nutrient solution delivered to the plants has to be increased. During the summer months, under continuous periods of intense light, the plants may not have recovered their water status overnight even when the daily over-drain targets have been met. The plants begin the day a dark bluegreen in color, an indication that they are already under water stress, even though the day has just begun. Under these circumstances the overdrain

targets for the day could be increased, but there is the associated risk of overwatering and decreasing root health and performance. In these cases it is advisable to consider one or two night waterings, one at approximately 10:00 p.m. or one at 2:00 am or both. Usually the night watering events are the same length of time as the minimum watering duration applied during the day. Night watering can also help increase the rate of fruit development, but there is an associated risk of fruit splitting if too much water is applied at night. The night waterings should not be continued indefinitely and the decision to use night watering events and to continue with night watering has to be based on the assessed needs of the crop. The management of the feed solution, and its delivery to the crop has to be relatively flexible to meet the changing needs of the crop. With experience, growers gain more confidence and skill in meeting and anticipating the changing needs of the crop throughout the crop cycle and through periods of fluctuating light levels. The general information presented in this section serves as a starting point and by following the principles of over-drain management, E.C. and pH monitoring and correction, a successful strategy for delivery of water and nutrients can be established. As with many things there is no one “right” way to apply water and nutrients to the crop. The use of leaching, although ensuring that salt levels do not accumulate to high levels in the root zone, does result in some “waste” of fertilizer solution as runoff. There are strategies that can be employed to minimize the waste associated with leaching. Collection and recirculation of the leachate, with an associated partial sterilization, or biofiltration of the nutrient solution is one approach. The sterilization or biofiltration steps are required in order to minimize the disease risk associated with recycling nutrient solutions. Some estimates place the fertilizer cost savings at between 30 to 40% when recirculation is used. In addition to being economical, recycling nutrient solutions is an environmentally sound practice. There is a limit to how long nutrient solutions can be recirculated, prolonged recycling of the same solution can negatively affect growth and yield. This is primarily associated with the accumulation of sulfate ions in the solution. In addition to sulfates, chlorides and bicarbonates also have a tendency to accumulate and can influence crop growth. The progressive accumulation of sulfates in recirculating solutions require occasional “refreshing” of the solution whereby the solution would have to be allowed to

leave the greenhouse as waste.

References Abou-Hadid, A.F., M.Z El-Shinawy, A.S. El-Beltagy, and S.W. Burrage. 1992. Relation between water use efficiency of sweet pepper grown under nutrient film technique and rockwool under protected cultivation. Acta Horticulturae 323:89-95. Adams, P. and L.C. Ho. 1995. Differential effects of salinity and humidity on growth and Ca status of tomato and cucumber grown in hydroponic culture. Acta Horticulturae 401:357-363. Anonymous. 1973. Growelectric Handbook 2; Lighting in greenhouses. The Electricity Council. London England. Bakker, J.C. 1989. The effects of temperature on flowering, fruit set and fruit development of glasshouse sweet pepper (Capsicum annuum L.). Journal of Horticultural Science 64(3):313-320. Blom, T.J. 1998. Air pollution in greenhouses. Regulatory Horticulture 24:15. Boivin, C., A. Gosselin, and M. J. Trudel. 1987. Effect of supplementary lighting on transplant growth and yield of greenhouse tomato. HortScience 22(6):1266-1268. Brugger, M.F., T.H. Short and W.L. Bauerle. 1987. An evaluation of horizontal air flow in six commercial greenhouses. American Society of Agricultural Engineers1987 summer meeting presentation number 374020. Chang, J. 1996. Screening greenhouse vents for insect pest control. Alberta Agriculture, Food and Rural Development Engineering Services Publication. De Koning, A.N.M. 1989. Development and growth of a commercially grown tomato crop. Acta Horticulturae 260:267-270. Demers, D.A., J. Charbonneau, J. and A Gosselin. 1991. Effects of supplementary lighting on the growth and productivity of greenhouse sweet pepper. Can J. Plant. Sci. 71:587-594. Demers, D.A., and A. Gosselin. 1998. Effects of supplemental light duration

on greenhouse sweet pepper plants and fruit yields. J. Amer. Soc Hort. Sci.123(2):202-207. Esiyok, D., E. Ozzambak and B. Eser. 1994. The effects of stem pruning on the yield and earliness of greenhouse peppers (Capsicum annum L. grossum cv.Kandil and 11B-14). Acta Horticulturae 366:293-300. Fierro, A., N. Tremblay and A. Gosselin. 1994. Supplemental carbon dioxide and light improved tomato and pepper seedling growth and yield. HortScience29(3):152-154. Fricke, A. and H. Krug. 1997. Influence of humidification and dehumidification on greenhouse climate as well as water relations and productivity of cucumber II. Influences on plants. Gartenbauwissenshaft 62(6):241-248. Gagne, S.L. Dehbi, D. Le Quere, F. Cayer, J.L. Morin, R. Lemay and N. Fournier. 1993. Increase of greenhouse tomato fruit yields by plant growth promoting rhizobacteria (PGPR) inoculated into the peat-based growing media. Soil Biol. Biochem. 25(2):269-272. Gauthier, L. 1992. GX: A small talk-based platform for greenhouse environmental control. Part I. Modeling and managing the physical system. Transactions of the ASAE 35(6):2003-2009. Hansen, C. W., J. Lynch and C. O. Ottosen. 1998. Response to phosphorus availability during vegetative and reproductive growth of chrysanthemum: I. Whole-plant carbon dioxide exchange. J. Amer. Soc. Hort. Sci. 132 (2):215-222. Hardgrove, M. R. 1992. Recirculation systems for greenhouse vegetables. Acta Horticulturae 303:85-92. Horbowicz, M, and A. Stepowska. 1995. Effect of growing conditions at greenhouse on vitamin E content in sweet pepper (Capsicum annuum L.) fruits. Acta Agrobotanica 48:61-67. Howard, R.J., J.A. Garland and W.L. Seaman. 1994. Diseases and pests of vegetable crops in Canada. The Canadian Phytopathological Society and the Entomological Society of Canada. Jarvis, W.R. 1992. Managing Diseases in Greenhouse Crops. APS Press, St. Paul, Minnesota.

Jespersen, L. M. and J. Willumsen. 1993. Production of compost in a heat composting plant and test of compost mixtures as a growing media for greenhouse cultures. Acta Horticulturae 342:127-142. Jolliet, O., B.J. Bailey, D.J. Hand, and K. Cockshull. 1993. Tomato yield in greenhouses related to humidity and transpiration. Acta Horticulturae 328:115-124. Jones, J.B. Jr., 1998. Plant Nutrition Manual. CRC Press, New York. Jones, J.W., E. Dayan, P. Jones, Y Hwang, and B. Jacobson. 1988. Modeling tomato growth for greenhouse environment control. American Society of Agricultural Engineers December 1988 winter meeting presentation number 88-7501. Jones, J.W., E. Dayan, H. Van Keulan, and H. Challa. 1989. Modeling tomato growth for optimizing greenhouse temperatures and carbon dioxide concentrations.Acta Horticulturae 248: 285-294. Jones, J.W., E. Dayan, L. H. Allen, H. Van Keulen, H. Challa. 1991. A dynamic tomato growth and yield model (TOMGRO). Transactions of the ASAE 34(2):663-672. Khah, E.M. and H.C. Passam. 1992. Flowering, fruit set and development of the fruit and seed of sweet pepper (Capsicum annuum L.) cultivated under conditions of high ambient temperature. Journal of Horticultural Science 67(2)251-258. Khosla, S. 1999. In “Greenhouse Vegetable Experts Discuss the Future of the Industry”, Tomato Magazine, August, 1999. Yakima Washington. Koppert Biological Systems 1999 “Bio-Plus” Technical Information Bulletin Lange, D. and H.J. Tantau. 1996. Climate management for disease control investigations on control strategies, plant densities and irrigation systems. Acta Horticulturae 406:105-113. LI-COR Inc. Radiation Measurement - LI COR Technical Bulletin. Lin, W.C., J.W. Hall, and M.E. Saltveit Jr. 1993. Ripening stage affects the chilling sensitivity of greenhouse-grown peppers. J. Amer. Soc. Hort. Sci. 118(6):791-795. Lin, W.C. and G.S. Block. 1997. Determination of water vapor in a small air sample by a non-dispersive infrared gas analyzer. HortScience 32(2):278-

281. Morard, P., A. Pujos, A. Bernadac, and G. Bertoni. 1996. Effect of temporary calcium deficiency on tomato growth and mineral nutrition. Journal of Plant Nutrition19(1):115-127. Nederhoff, E.M. 1994. Effects of CO2 concentration on photosynthesis, transpiration and production of greenhouse fruit vegetable crops. University of Wageningen, The Netherlands. Nederhoff, E. M. and J. G. Vegter. 1994. Canopy photosynthesis of tomato, cucumber and sweet pepper in greenhouses: measurements compared to models.Annals of Botany 73: pp 412 - 427. Ng, K. and T. van der Gulick, 1999. Bio-Sand Filtration. British Columbia Ministry of Agriculture and Food Factsheet 99-01. Padem H. and R. Alan. 1994. The effects of some substrates on yield and chemical composition of pepper under greenhouse conditions. Acta Horticulturae 366: 445 - 451. Papadakis, G., A. Frangoudakis, and S. Kyritsis. 1994. Experimental investigation and modelling of heat and mass transfer between a tomato crop and the greenhouse environment. J. Agric. Engng Res. 57:217-227. Papadopoulos, A.P., and S. Pararajasingham. 1997. The influence of plant spacing on light interception and use in greenhouse tomato (Lycopersicon esculentum Mill.): A review. Scientia Horticulturae 69:1-27. Pfadt, R.E. ed. 1978. Fundamentals of Applied Entomology 3rd edition. Macmillan Publishing Co., Inc. New York. Portree, J. 1996. Greenhouse vegetable production guide for commercial growers. Province of British Columbia Ministry of Agriculture, Fisheries and Food. Pressman, E., H. Moshkovitch, K. Rosenfeld, R. Shaked, B. Gamliel and B. Aloni. 1998. Influence of low night temperatures on sweet pepper flower quality and the effect of repeated pollinations, with viable pollen, on fruit setting. Journal of Horticultural Science & Biotechnology 73(1)131-136. Pulupol, L.U., M.H. Behboudian, and K.J. Fisher. 1996. Growth, yield and postharvest attributes produced under deficit irrigation. HortScience 31(6):926-929.

Rijkdijk, A.A. and G. Houter. 1993. Validation of a model for energy consumption, CO2consumption and crop production (ECP-model). Acta Horticulturae328:125-131. Rodov, V., S. Ben-Yehoshua, T. Fierman and D. Fang. 1995. Modifiedhumidity packaging reduces decay of harvested red bell pepper fruit. HortScience 30(2):299-302. Romero-Aranda, R. and J.J. Longuenesse. 1995. Modelling the effect of air vapour pressure deficit on leaf photosynthesis of greenhouse tomatoes: The importance of leaf conductance to CO2. Journal of Horticultural Science 70(3):423-432. Schon, M.K. 1993. Effects of foliar antitranspirant or calcium nitrate applications on yield and blossom-end rot occurrence in greenhousegrown peppers.Journal of Plant Nutrition 16(6):1137-1149. Seginer, I. and R.W. McClendon. 1992. Methods for optimal control of the greenhouse environment. Transactions of the ASAE 35(4)1299-1307. Seginer, I. 1996. Optimal control of the greenhouse environment: an overview. Acta Horticulturae 406:191-201. Seginer, I., Y. Hwang, T. Boulard, J.W. Jones. 1996. Mimicking an expert greenhouse grower with a neural-net policy. Transactions of the ASAE 39(1):299-306. Shina, G. and I. Seginer. 1989. Optimal management of tomato growth in greenhouses. Acta Horticulturae 248:307-313. Simon, L., T.J. Smalley, J. Benton Jones Jr., and F.T. Lasseigne. 1994. Aluminum toxicity in tomato. Part 1. Growth and mineral nutrition. Journal of Plant Nutrition17(2&3):293-306. Simon, L., M. Kieger, S.S. Sung, and T. J. Smalley. 1994. Aluminum toxicity in tomato. Part 2. Leaf gas exchange, chlorophyll content, and invertase activity.Journal of Plant Nutrition 17(2&3):307-317. Slack, G., J.S. Fenlon and D.W. Hand. 1988. The effects of summer CO2 enrichment and ventilation temperatures on the yield, quality and value of glasshouse tomatoes. Journal of Horticultural Science 63(1):119-129. Stanghellini, C., W.Th.M. Van Meurs. 1992. Environmental control of greenhouse crop transpiration. J. Agric. Engng Res. 51:297-311.

Styer, R.C. and D.S. Koranski. 1997. Plug and transplant production, a grower’s guide. Ball Publishing, Batavia, Illinois. USA. Tilley, D.E. 1979. Contemporary College Physics. Benjamin/Cummings Publishing Company. Menlo Park, California. Tootil, E. and S. Blackmore. 1984. The Facts on File Dictionary of Botany. Market House Books Ltd. Aylesbury, U. K. Tremblay, N. and A. Gosselin. 1998. Effect of carbon dioxide enrichment and light. HortTechnology 8(4):524-528. Van Meurs, W.Th.M., and C. Stanghellini. 1992. Environmental control of a tomato crop using a transpiration model. Acta Horticulturae 303:23-30. Weiler, T.C. and M. Sailus. 1996. Water and nutrient management for greenhouses. Northeast Regional Agricultural Engineering Service Cooperative Extension. Ithaca, New York, USA. Whaley-Emmons, C.L., and J.W. Scott. 1997. Environmental and physiological effects on cuticle cracking in tomato. J. Amer. Soc. Hort. Sci. 122(6):797-801. Wilson, C.L., and W.E. Loomis. 1967. Botany 4th edition. Holt, Rinehart and Winston. New York, USA. Wilson, J.W., D.W. Hand and M.A. Hannah. 1992. Light interception and photosynthetic efficiency in some glasshouse crops. Journal of Experimental Botany43(248):363-373. Wittwer, S.H. and S. Honma. 1979. Greenhouse tomatoes, lettuce and cucumbers. Michigan State University Press. East Lansing, USA. Wolfe, E.W., D.T. Topoleski, N.A. Gundersheim and B.A. Ingall. 1995. Growth and yield sensitivity of four vegetable crops to soil compaction. J. Amer. Soc. Hort. Sci. 120(6):956-963. Zabri, A.W., and S.W. Burrage. 1997. The effects of vapour pressure deficit (VPD) and enrichment with CO2 on water relations, photosynthesis, stomatal conductance and plant growth of sweet pepper (Capsicum annum L.) grown by NFT. Acta Horticulturae 449(2):561-567. Zekki, H., L. Gauthier and A. Gosselin. 1996. Growth, productivity, and mineral composition of hydroponically cultivated greenhouse tomatoes, with or without nutrient solution recycling. J. Amer. Soc. Hort. Sci.

121(6):1082-1088.

Bibliography Abou-Hadid, A.F., M.Z El-Shinawy, A.S. El-Beltagy, and S.W. Burrage. 1992. Relation between water use efficiency of sweet pepper grown under nutrient film technique and rockwool under protected cultivation. Acta Horticulturae 323: 89-95. Adams, P. and L.C. Ho. 1995. Differential effects of salinity and humidity on growth and Ca status of tomato and cucumber grown in hydroponic culture. Acta Horticulturae 401: 357-363. Ahmad, Shahid (2013). Hill Agriculture. Astral Publishing house , New Delhi pp550. Ahmad, Shahid (2009). Plant Disease Management for Sustainable Agriculture Daya Publishing house New Delhi. pp373. Ahmad, Shahid (2012). Ali Anwar and P.K. Sharma (2011). Plant Disease Management on Horticultural Crops, Daya Publishing House New Delhi. pp 405. Ahmad, Shahid (2012). Recent Trends in Plant Diseases Management in India, Kalyani Publisher, Ludhiana, India. pp 478. Ahmad, Shahid and Ali Anwar (2014). Terminology on Plant Pathology, Jaya Publishing House, Delhi-110006, pp. 159. Ahamad, Shahid and Udit Narain (2007) Eco-friendly Management of Plant Diseases, Daya Publishing House New Delhi. pp 412. Anonymous. 1973. Growelectric Handbook 2; Lighting in greenhouses. The Electricity Council. London England. Aylsworth, Jean. 1993. Biological controls catch on with growers. Greenhouse Grower. December. pp. 77-78, 80-81. Baker, J. R. 1994. Insects and Related Pests of Flowers and Foliage Plants. Some important, common, and potential pests in the southeastern United

States. North Carolina Cooperative Extension Service. 105 p. Bakker, J.C. 1989. The effects of temperature on flowering, fruit set and fruit development of glasshouse sweet pepper (Capsicum annuum L.). Journal of Horticultural Science 64(3): 313-320. Blom, T.J. 1998. Air pollution in greenhouses. Regulatory Horticulture 24: 15. Boivin, C., A. Gosselin, and M. J. Trudel. 1987. Effect of supplementary lighting on transplant growth and yield of greenhouse tomato. HortScience 22(6): 1266-1268. Boucher, T. J. Cole Crop “Worms”. Caterpillar Pests. UCONN IPM Factsheet. Boucher, T. J. and R. G. Adams. 1993. Integrated Pest Management Guide for Cole Crops. UConn Coop. Ext. Syst. Pub. 93-19, 21 p. Brugger, M.F., T.H. Short and W.L. Bauerle. 1987. An evaluation of horizontal air flow in six commercial greenhouses. American Society of Agricultural Engineers1987 summer meeting presentation number 374020. Capinera, J. 2009. Saltmarsh Caterpillar, Estigmene acrea (Dury) (Insecta: Lepidoptera: Arctiidae). Casey, C. (Ed.) 2000. Integrated Pest Management for Bedding Plants. A Scouting and Pest Management Guide. New York State IPM Program. IPM Bulletin No. 407. 117 p. Casey, C. 2000. Integrated Pest Management for Bedding Plants. A Scouting and Pest Management Guide. Cornell Cooperative Extension Publication No. 407. 117 pp. Casey, C. Ed. 1999. Integrated Pest Management for Bedding Plants. A Scouting and Pest Management Guide. Cornell Cooperative Extension Pub. No. 407, 109 p. Catlin, N. 2012. Two-Spotted Spider Mites and Edema on Geranium. E-Gro Alter, May 2, 2012. Chang, J. 1996. Screening greenhouse vents for insect pest control. Alberta Agriculture, Food and Rural Development Engineering Services Publication.

Chase, A.R. 1998. New bactericides and fungicides for disease control on ornamentals. Greenhouse Product News. December. pp. 22-24. Cloyd, R. 2011. Twospotted Spider Mite Management in Greenhouses. Floribytes. March 2011. Cloyd, R. 2012. Plant Health: Combating Caterpillars. Greenhouse Management. Cloyd, R. 2015. Ecology of Fungus Gnats (Bradysia spp.) in Greenhouse Production Systems Associated with Disease-Interactions and Alternative Management Strategies. Insects. 6: 325-332. Cox D. 2014. Organic Fertilizers - Thoughts on Using Liquid Organic Fertilizers for Greenhouse Plants. Sept.-Oct. Floral Notes 27(2). Cox D. 2016 Plant Response to Nature’s Source and EcoVita Organic Fertilizers vs Plantex Chemical Fertilizer. July-Aug. Floral Notes 29(1). Cox, D. and L. Craker. 1993. Growing Herbs as Bedding Plants. University of Massachusetts Floral Notes. 6(3): 2-6. Cranshaw, W. 2004. Garden Insects of North America. Princeton University Press. Princeton, NJ. 656 p. De Koning, A.N.M. 1989. Development and growth of a commercially grown tomato crop. Acta Horticulturae 260: 267-270. Demers, D.A., and A. Gosselin. 1998. Effects of supplemental light duration on greenhouse sweet pepper plants and fruit yields. J. Amer. Soc Hort. Sci.123(2): 202-207. Demers, D.A., J. Charbonneau, J. and A Gosselin. 1991. Effects of supplementary lighting on the growth and productivity of greenhouse sweet pepper. Can J. Plant. Sci. 71: 587-594. Dutky, Ethel. 1995. Here’s how to cut your losses due to disease. GMPro. October. p. 63-65. Esiyok, D., E. Ozzambak and B. Eser. 1994. The effects of stem pruning on the yield and earliness of greenhouse peppers (Capsicum annum L. grossum cv. Kandil and 11B-14). Acta Horticulturae 366: 293-300. Ferguson, G., G. Murphy, and L. Shipp. 2014. Fungus Gnats and Shoreflies in Greenhouse Crops. Ontario Ministry of Agriculture and Food Fact Sheet 14-003.

Fierro, A., N. Tremblay and A. Gosselin. 1994. Supplemental carbon dioxide and light improved tomato and pepper seedling growth and yield. HortScience 29(3): 152-154. Fricke, A. and H. Krug. 1997. Influence of humidification and dehumidification on greenhouse climate as well as water relations and productivity of cucumber II. Influences on plants. Gartenbauwissenshaft 62(6): 241-248. Gagne, S.L. Dehbi, D. Le Quere, F. Cayer, J.L. Morin, R. Lemay and N. Fournier. 1993. Increase of greenhouse tomato fruit yields by plant growth promoting rhizobacteria (PGPR) inoculated into the peat-based growing media. Soil Biol. Biochem. 25(2): 269-272. Gamliel, A. et al. No date. Solarization for the Recycling of Container Media. The Hebrew University of Jerusalem, Rehovot, Israel. Unpublished manuscript. 8 p. Garibaldi, Angelo, and M. Lodovica Bullino. 1991. Soil solarization in Southern European countries, with emphasis on soilborne disease control of protected crops. pp. 227-235. In: Jaacov Katan and James E. DeVay (ed.) Soil Solarization. CRC Press, Boca Raton, FL. Gauthier, L. 1992. GX: A small talk-based platform for greenhouse environmental control. Part I. Modeling and managing the physical system. Transactions of the ASAE 35(6): 2003-2009. Gentile, A.G., and D.T. Scanlon; Revised by Tina Smith. 1992. A Guide to Insects and Related Pests of Floricultural Crops in New England: For Commercial Growers. University of Massachusetts Cooperative Extension System. 36 p. Giblin, R.M., and S.D. Verkade. 1987. Solarization of small volumes of potting soil for disinfection of plant-parasitic nematodes. p. 174-176. In: Proc. Fla. State Hort. Soc. Vol. 100. Gill, S and J. Sanderson. 1998. Ball Identification Guide to Greenhouse Pests and Beneficals. Ball Publishing. Batavia, Ill. 244 p. Gill, S. 2013. Greenhouse IPM Pest Alert: European Pepper Moth. University of Maryland Gill, S. and J. Sanderson. 1998. Ball Identification Guide to Greenhouse Pests and Beneficials. Ball Publishing. Batavia, Ill. 244 p.

Gill, S., C. Casey, M. Raupp, . Davidson. 1993. The Whitefly Problem in Maryland Greenhouses. Cooperative Extension Service, University of Maryland. IPM Factsheet. 4p. Gilrein, D. 2001 New Miticides for Greenhouse, Nursery and Professional Landscape Use. Plugged In. The Newsletter of the Connecticut Greenhouse Growers Association. Issue 1. pp. 4-7. Gilrein, D. G. 2015. Time to Think About Aphids - Again. E-Gro Alert. 4(8) February 2015. Gindrat, D. 1979. Biological soil disinfection. p. 253.287. In: D. Mulder (ed.) Soil Disinfection. Elsevier Scientific Publishing Co., New York, NY. Hansen, C. W., J. Lynch and C. O. Ottosen. 1998. Response to phosphorus availability during vegetative and reproductive growth of chrysanthemum: I. Whole-plant carbon dioxide exchange. J. Amer. Soc. Hort. Sci. 132 (2): 215-222. Hardgrove, M. R. 1992. Recirculation systems for greenhouse vegetables. Acta Horticulturae 303: 85-92. Hoddle, M. 1994. Encarsia formosa: A parasitic wasp that attacks whiteflies. UMass Floral Notes 6(5): 5-7. Hoddle, M., R. Van Driesche, S. Roy, T. Smith, P. Lopes & J. Sanderson. 1996. A Grower’s Guide to Using Biological Control for Silverleaf Whitefly on Poinsettias in the Northeast United States. University of Massachusetts Cooperative Extension System Floral Facts. 4p. Hoffman, M. and J. Sanderson. 1993. Melon Aphid. Cornell Cooperative Extension Factsheet. No. 750.50 1 p. Hoitink, H.A.J., Y. Inbar, and M.J. Boehm. 1991. Status of compost-amended potting mixes naturally suppressive to soil-borne diseases of floricultural crops. Plant Disease. September. pp. 869-873. Hoitink, Harry A., and Peter C. Fahy. 1986. Basis for the control of soilborne plant pathogens with composts. Annual Reviews of Phytopathology. Vol. 24. pp. 93-114. Horbowicz, M, and A. Stepowska. 1995. Effect of growing conditions at greenhouse on vitamin E content in sweet pepper (Capsicum annuum L.) fruits. Acta Agrobotanica 48: 61-67.

Horiuchi, Seizo. 1991. Soil solarization in Japan. pp. 215, 218-223, 225. In: Jaacov Katan and James E. DeVay (ed.) Soil Solarization. CRC Press, Boca Raton, FL. Howard, R., J., J.A. Garland, and W.L. Seaman (Ed). 1994. Herbs and Spices.In Diseases and Pests of Vegetable Crops in Canada. The Canadian Phytopathological Society and Entomological Society of Canada. Ottawa, Canada. Howard, R.J., J.A. Garland and W.L. Seaman. 1994. Diseases and pests of vegetable crops in Canada. The Canadian Phytopathological Society and the Entomological Society of Canada. Jandricic, S. and J. Sanderson, 2011. Early Season Pest Threat. Greenhouse Canada, 12-14. Jarvis, W.R. 1992. Managing Diseases in Greenhouse Crops. APS Press, St. Paul, Minnesota. Jespersen, L. M. and J. Willumsen. 1993. Production of compost in a heat composting plant and test of compost mixtures as a growing media for greenhouse cultures. Acta Horticulturae 342: 127-142. Jolliet, O., B.J. Bailey, D.J. Hand, and K. Cockshull. 1993. Tomato yield in greenhouses related to humidity and transpiration. Acta Horticulturae 328: 115-124. Jones, J.B. Jr., 1998. Plant Nutrition Manual. CRC Press, New York. Jones, J.W., E. Dayan, H. Van Keulan, and H. Challa. 1989. Modeling tomato growth for optimizing greenhouse temperatures and carbon dioxide concentrations. Acta Horticulturae 248: 285-294. Jones, J.W., E. Dayan, L. H. Allen, H. Van Keulen, H. Challa. 1991. A dynamic tomato growth and yield model (TOMGRO). Transactions of the ASAE 34(2): 663-672. Jones, J.W., E. Dayan, P. Jones, Y Hwang, and B. Jacobson. 1988. Modeling tomato growth for greenhouse environment control. American Society of Agricultural Engineers December 1988 winter meeting presentation number 88-7501. Khah, E.M. and H.C. Passam. 1992. Flowering, fruit set and development of the fruit and seed of sweet pepper (Capsicum annuum L.) cultivated under

conditions of high ambient temperature. Journal of Horticultural Science 67(2): 251-258. Khosla, S. 1999. In “Greenhouse Vegetable Experts Discuss the Future of the Industry”, Tomato Magazine, August, 1999. Yakima Washington. Klassen, Parry. 1993. Mulling over methyl bromide. Greenhouse Grower. August. p. 118 & 120. Koppert Biological Systems 1999 “Bio-Plus” Technical Information Bulletin Kuack, David. 1995. Janet Bandy on implementing an effective IPM program. Greenhouse Management and Production. April. pp. 56-57. Lal, B., Shahid Ahmad and De (2016). Modeling in Communication behaviors of Farmers, Published by Astral International Publisher, New Delhi. Lamb, E., B. Eshenaur, N. Mattson, and J. Sanderson. 2013. Practical Suggestions for Managing Fungus Gnats in the Greenhouse. Cornell University Ornamental IPM Factsheet. Lange, D. and H.J. Tantau. 1996. Climate management for disease control investigations on control strategies, plant densities and irrigation systems. Acta Horticulturae 406: 105-113. LI-COR Inc. Radiation Measurement - LI COR Technical Bulletin. Lin, W.C. and G.S. Block. 1997. Determination of water vapor in a small air sample by a non-dispersive infrared gas analyzer. HortScience 32(2): 278-281. Lin, W.C., J.W. Hall, and M.E. Saltveit Jr. 1993. Ripening stage affects the chilling sensitivity of greenhouse-grown peppers. J. Amer. Soc. Hort. Sci. 118(6): 791-795. Lindquist, Richard K. 1998. Evaluations of non-conventional pesticides for insect and mite control on greenhouse ornamental plants. Greenhouse Product News. July. pp. 52-55. Mahrer, Yitzhak. 1991. Physical properties of solar heating of soils by plastic mulching in the field and in glasshouses and simulation models. pp. 75, 81-86. In: Jaacov Katan James E. DeVay (ed.) Soil Solarization. CRC Press, Boca Raton, FL. Malais, M. And W.J. Ravensburg. 2003. Knowing and Recognizing: The

biology of glasshouse pests and their natural enemies. Koppert Biological Systems. 288 pp. Mattson N. Substrates and Fertilizers for Organic Vegetable Transplant Production. Cornell Greenhouse Horticulture, Cornell University. McKenzie, C. L., J. A. Bethke, F.J. Byrne, J. R. Chamberlin, T.J. Dennehy, A. M. Dickey, D.Gilrein, P.M. Hall, S.Ludwig, R.D. Oetting, L.S. Osborne, and Shatters, R.G. 2012. Distribution of Bemisia tabaci (Hemiptera: Aleyrodidae) Biotypes in North America After the Q Invasion. Journal of Economic Entomology 105(3): 753-766. Meers, T. and R. Cloyd. 2005. Egg-Laying Preference of Female Fungus Gnat Bradysia sp. nr. coprophila (Diptera: Sciaridae) on Three Different Soilless Substrates. Journal of Economic Entomology. 98(6): 1937-1942. Morard, P., A. Pujos, A. Bernadac, and G. Bertoni. 1996. Effect of temporary calcium deficiency on tomato growth and mineral nutrition. Journal of Plant Nutrition 19(1): 115-127. Nederhoff, E. M. and J. G. Vegter. 1994. Canopy photosynthesis of tomato, cucumber and sweet pepper in greenhouses: measurements compared to models. Annals of Botany 73: pp 412 - 427. Nederhoff, E.M. 1994. Effects of CO2 concentration on photosynthesis, transpiration and production of greenhouse fruit vegetable crops. University of Wageningen, The Netherlands. Ng, K. and T. van der Gulick, 1999. Bio-Sand Filtration. British Columbia Ministry of Agriculture and Food Factsheet 99-01. Oetting, R. and J. Chong. 2002. Spider Mites. In Proceedings of the Society of American Florists’ 18th Annual Conference on Insect and Disease Management on Ornamentals. San Diego, CA. Organic Greenhouse Vegetable Production, Potting Mixes for Certified Organic Production, Organic Greenhouse Tomato Production, Plug and Transplant Production for Organic Systems, ATTRA - National Sustainable Agriculture Information Service. Padem H. and R. Alan. 1994. The effects of some substrates on yield and chemical composition of pepper under greenhouse conditions. Acta Horticulturae 366: 445-451.

Papadakis, G., A. Frangoudakis, and S. Kyritsis. 1994. Experimental investigation and modelling of heat and mass transfer between a tomato crop and the greenhouse environment. J. Agric. Engng Res. 57: 217-227. Papadopoulos, A.P., and S. Pararajasingham. 1997. The influence of plant spacing on light interception and use in greenhouse tomato (Lycopersicon esculentum Mill.): A review. Scientia Horticulturae 69: 1-27. Pfadt, R.E. ed. 1978. Fundamentals of Applied Entomology 3rd edition. Macmillan Publishing Co., Inc. New York. Portree, J. 1996. Greenhouse vegetable production guide for commercial growers. Province of British Columbia Ministry of Agriculture, Fisheries and Food. Pressman, E., H. Moshkovitch, K. Rosenfeld, R. Shaked, B. Gamliel and B. Aloni. 1998. Influence of low night temperatures on sweet pepper flower quality and the effect of repeated pollinations, with viable pollen, on fruit setting. Journal of Horticultural Science & Biotechnology 73(1): 131136. Pulupol, L.U., M.H. Behboudian, and K.J. Fisher. 1996. Growth, yield and postharvest attributes produced under deficit irrigation. HortScience 31(6): 926-929. Reuveni, R., N. Dudai, and E. Putievski. 1998. Nufar: A Sweet Basil Cultivar Resistant to Fusarium Wilt. HortScience. 33(1): 159. Richter, C. 1999. Success with Mints. GrowerTalks. 63 (1): 105,110. Rijkdijk, A.A. and G. Houter. 1993. Validation of a model for energy consumption, CO2 consumption and crop production (ECP-model). Acta Horticulturae 328: 125-131. Rodov, V., S. Ben-Yehoshua, T. Fierman and D. Fang. 1995. Modifiedhumidity packaging reduces decay of harvested red bell pepper fruit. HortScience 30(2): 299-302. Romero-Aranda, R. and J.J. Longuenesse. 1995. Modelling the effect of air vapour pressure deficit on leaf photosynthesis of greenhouse tomatoes: The importance of leaf conductance to CO2. Journal of Horticultural Science 70(3): 423-432. Sanderson, J. and S. Jandricic. 2016. Out-foxing the Foxglove Aphid.

GrowerTalks. October 28, 2016. Sanderson, J.P. 1995. Total whitefly identification and control. GrowerTalks. 59(6): 32-44. Schon, M.K. 1993. Effects of foliar antitranspirant or calcium nitrate applications on yield and blossom-end rot occurrence in greenhousegrown peppers.Journal of Plant Nutrition 16(6): 1137-1149. Seginer, I. and R.W. McClendon. 1992. Methods for optimal control of the greenhouse environment. Transactions of the ASAE 35(4): 1299-1307. Seginer, I. 1996. Optimal control of the greenhouse environment: an overview. Acta Horticulturae 406: 191-201. Seginer, I., Y. Hwang, T. Boulard, J.W. Jones. 1996. Mimicking an expert greenhouse grower with a neural-net policy. Transactions of the ASAE 39(1): 299-306. Shina, G. and I. Seginer. 1989. Optimal management of tomato growth in greenhouses. Acta Horticulturae 248: 307-313. Shore, S. 2003. Growing and Selling Fresh-Cut Herbs. Ball Publishing, Batavia, IL. 483 p. Thomas, P. 1997. The Challenges. Simon, L., M. Kieger, S.S. Sung, and T. J. Smalley. 1994. Aluminum toxicity in tomato. Part 2. Leaf gas exchange, chlorophyll content, and invertase activity.Journal of Plant Nutrition 17(2&3): 307-317. Simon, L., T.J. Smalley, J. Benton Jones Jr., and F.T. Lasseigne. 1994. Aluminum toxicity in tomato. Part 1. Growth and mineral nutrition. Journal of Plant Nutrition17(2&3): 293-306. Slack, G., J.S. Fenlon and D.W. Hand. 1988. The effects of summer CO2 enrichment and ventilation temperatures on the yield, quality and value of glasshouse tomatoes. Journal of Horticultural Science 63(1): 119-129. Smith, T. and L. Pundt. 2001. Pest Management for Vegetable Bedding Plants. New England Greenhouse Conference Fact Sheet 1. 8 p. Stanghellini, C., W.Th. M. Van Meurs. 1992. Environmental control of greenhouse crop transpiration. J. Agric. Engng Res. 51: 297-311. Stocks, S. D. and A. Hodges. 2013. Featured Creatures: European pepper moth or southern European marsh pyralid. University of Florida

Publication EENY-508. Styer, R.C. and D.S. Koranski. 1997. Plug and transplant production, a grower’s guide. Ball Publishing, Batavia, Illinois. U.S.A. The National Organic Program http://www.ams.usda.gov/nop/NOP/Nophome.html

Guidelines:

Thomas, C. Greenhouse IPM with an Emphasis on Biocontrols. Publication No. AGRS-96. 89 p. Pennsylvania Integrated Pest Management Program. Tilley, D.E. 1979. Contemporary College Physics. Benjamin/Cummings Publishing Company. Menlo Park, California. Tootil, E. and S. Blackmore. 1984. The Facts on File Dictionary of Botany. Market House Books Ltd. Aylesbury, U. K. Tremblay, N. and A. Gosselin. 1998. Effect of carbon dioxide enrichment and light. HortTechnology 8(4): 524-528. Van Meurs, W.Th.M., and C. Stanghellini. 1992. Environmental control of a tomato crop using a transpiration model. Acta Horticulturae 303: 23-30. Weiler, T.C. and M. Sailus. 1996. Water and nutrient management for greenhouses. Northeast Regional Agricultural Engineering Service Cooperative Extension. Ithaca, New York, USA. Whaley-Emmons, C.L., and J.W. Scott. 1997. Environmental and physiological effects on cuticle cracking in tomato. J. Amer. Soc. Hort. Sci. 122(6): 797-801. Wilson, C.L., and W.E. Loomis. 1967. Botany, 4th edition. Holt, Rinehart and Winston. New York, USA. Wilson, J.W., D.W. Hand and M.A. Hannah. 1992. Light interception and photosynthetic efficiency in some glasshouse crops. Journal of Experimental Botany 43(248): 363-373. Wittwer, S.H. and S. Honma. 1979. Greenhouse tomatoes, lettuce and cucumbers. Michigan State University Press. East Lansing, USA. Wolfe, E.W., D.T. Topoleski, N.A. Gundersheim and B.A. Ingall. 1995. Growth and yield sensitivity of four vegetable crops to soil compaction. J. Amer. Soc. Hort. Sci. 120(6): 956-963. Zabri, A.W., and S.W. Burrage. 1997. The effects of vapour pressure deficit

(VPD) and enrichment with CO2 on water relations, photosynthesis, stomatal conductance and plant growth of sweet pepper (Capsicum annum L.) grown by NFT. Acta Horticulturae 449(2): 561-567. Zekki, H., L. Gauthier and A. Gosselin. 1996. Growth, productivity, and mineral composition of hydroponically cultivated greenhouse tomatoes, with or without nutrient solution recycling. J. Amer. Soc. Hort. Sci. 121(6): 1082-1088.