Practical Handbook on Soil Protists (Springer Protocols Handbooks) [2024 ed.] 1071637495, 9781071637494

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N. Amaresan Komal A. Chandarana  Editors

Practical Handbook on Soil Protists

SPRINGER PROTOCOLS HANDBOOKS

For further volumes: http://www.springer.com/series/8623

Springer Protocols Handbooks collects a diverse range of step-by-step laboratory methods and protocols from across the life and biomedical sciences. Each protocol is provided in the Springer Protocol format: readily-reproducible in a step-by-step fashion. Each protocol opens with an introductory overview, a list of the materials and reagents needed to complete the experiment, and is followed by a detailed procedure supported by a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. With a focus on large comprehensive protocol collections and an international authorship, Springer Protocols Handbooks are a valuable addition to the laboratory.

Practical Handbook on Soil Protists Edited by

N. Amaresan and Komal A. Chandarana C.G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India

Editors N. Amaresan C.G. Bhakta Institute of Biotechnology Uka Tarsadia University Surat, Gujarat, India

Komal A. Chandarana C.G. Bhakta Institute of Biotechnology Uka Tarsadia University Surat, Gujarat, India

ISSN 1949-2448 ISSN 1949-2456 (electronic) Springer Protocols Handbooks ISBN 978-1-0716-3749-4 ISBN 978-1-0716-3750-0 (eBook) https://doi.org/10.1007/978-1-0716-3750-0 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface Soil heterotrophic protists are a diverse group of eukaryotic organisms divided into four groups: naked amoebae, ciliates, flagellates, and testate amoebae. These bacterivorous and/or mycophagous protists impart their role by performing various ecological functions, such as consumers of the soil food web and nutrient cycling. However, this area of research is less focused due to methodological constraints. The success of protistological research and comparative studies depends on the successful isolation and identification of these microbes. The present manual covers a wide range of basic techniques associated with the observation, isolation, identification, and staining of these immensely important organisms. The introduction to each protocol explains its importance and application. A proper understanding and, sometimes, a slight modification in the protocol will help the researchers to achieve their goal. This book consists of six parts with 33 protocol chapters. Part I deals with the abundance and diversity calculations of soil protists. Parts II, III, and IV present the basic techniques for isolation, enumeration, and enrichment of the four groups of soil protists separately. Part V provides the number of different molecular and staining techniques used to identify the different protists from soil. Part VI describes the isolation, identification, and staining of the selective protistan group. This book will help postgraduate students, research scholars, and most importantly, inexperts in this area of protistological research studies. Some basic techniques, which are of utmost importance for separating, observing, and identifying this group of organisms, will fill the gaps in culturing these microbes in the laboratory. Surat, Gujarat, India

N. Amaresan Komal A. Chandarana

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

ABUNDANCE AND SPECIES RICHNESS CALCULATION OF PROTISTS FROM SOIL

1 Soil Sampling, Processing, and Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jean C. V. Dutra and Maria C. P. Batitucci 2 Initial Observation of Protist from Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Veysel Turan 3 Abundance Calculation of Naked and Testate Amoebae from Soil. . . . . . . . . . . . . Urjita Sheth 4 Abundance Calculation of Mycophagous Amoebae . . . . . . . . . . . . . . . . . . . . . . . . . Urjita Sheth 5 Abundance Calculation of Ciliates from Soil. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Urjita Sheth 6 Abundance Calculation of Flagellates from Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Urjita Sheth 7 Methodology to Study Species Diversity of Naked Amoeba and Testate Amoeba. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hetvi Naik, Sapna Chandwani, and Natarajan Amaresan 8 Non-flooded Petri Plate Method to Study the Species Diversity of Flagellates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hetvi Naik, Sapna Chandwani, and Natarajan Amaresan

PART II

11 12

3 7 13 23 29 33

37

43

ENUMERATION TECHNIQUES

9 Enumeration of Testate Amoeba Through Direct Count from Soil. . . . . . . . . . . . Komal A. Chandarana, Hetvi Naik, and Natarajan Amaresan 10 Most Probable Number (MPN) Method for Enumeration of Soil Protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jun Murase

PART III

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55

ISOLATION OF PROTIST FROM SOIL

Standard Solutions and Media Used for Isolation of Soil Protists . . . . . . . . . . . . . Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Isolation Techniques for Amoeba . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jun Murase

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63 69

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13 14

Contents

Isolation of Ciliates and Flagellates from Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amol D. Theng, Bhavana V. Mohite, and Satish V. Patil Isolation and Enumeration of Mycophagous Protist. . . . . . . . . . . . . . . . . . . . . . . . . Amol D. Theng, Bhavana V. Mohite, and Satish V. Patil

PART IV 15 16 17 18 19 20

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ENRICHMENT OF SOIL PROTISTS ON LABORATORY MEDIA

Media Used for Enrichment of Soil Protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Komal A. Chandarana and Natarajan Amaresan Enrichment of Naked Amoebae Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Enrichment of Ciliates and Flagellates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Enrichment of Mycophagous Protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Mono-axenic Cultivation of Protists. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Komal A. Chandarana and Natarajan Amaresan Axenic Cultivation of Soil Protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan

PART V

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89 95 101 107 111 117

IDENTIFICATION TECHNIQUES FOR SOIL PROTISTS

Classical Wet Mount Method for Observing Live Protists. . . . . . . . . . . . . . . . . . . . Komal A. Chandarana and Natarajan Amaresan 22 SEM: Sample Preparation, Fixation, and Staining of Protists . . . . . . . . . . . . . . . . . Shraddha Saha 23 TEM: Sample Preparation, Fixation, and Staining of Protists . . . . . . . . . . . . . . . . . Shraddha Saha 24 Protargol Staining Technique for Identification of Soil Protists . . . . . . . . . . . . . . . Mehreen Shah and Sirajuddin Ahmed 25 FLUTAX Staining of Cilliates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Swati Patel and Vimal Prajapati 26 DNA Extraction from Trophozoites and Cysts Using Manual Protocol. . . . . . . . Kejal Gohil, Komal A. Chandarana, G. Jawahar, and Natarajan Amaresan 27 DNA Isolation from Single Ciliate Isolate. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Swati Patel and Vimal Prajapati 28 Species Identification Through Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vimalkumar Prajapati, Swati Patel, Vaibhavkumar Mehta, and B. Z. Dholakiya 29 DNA Barcoding Techniques for Protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amit Gamit and Dhruti Amin

125 127 133 137 143 147

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Contents

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Unequivocal Identification Through Fluorescence In Situ Hybridization (FISH) Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 Jean C. V. Dutra and Maria C. P. Batitucci

PART VI

OTHER PROTISTAN

31

Isolation, DNA Extraction, Amplification, and Gel Electrophoresis of Single-Celled Nonmarine Foraminifera (Rhizaria) . . . . . . . . . . . . . . . . . . . . . . . . 181 Maria Holzmann 32 Isolation and Cultivation and Staining of Paramecium . . . . . . . . . . . . . . . . . . . . . . 189 Satish V. Patil, Sunil H. Koli, Bhavana V. Mohite, Jitendra D. Salunkhe, Amol D. Theng, and Atharv S. Patil 33 Isolation and Cultivation of Euglena . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 Satish V. Patil, Sunil H. Koli, Bhavana V. Mohite, Narendra S. Salunkhe, and Atharv S. Patil Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

201

Contributors SIRAJUDDIN AHMED • Department of Environmental Science and Engineering, Jamia Millia Islamia (Central University), New Delhi, India NATARAJAN AMARESAN • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India DHRUTI AMIN • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India MARIA C. P. BATITUCCI • Laboratorio de Gene´tica Vegetal e Toxicologica, Departamento de Cieˆncias Biologicas, Universidade Federal do Espı´rito Santo, Vitoria, Espı´rito Santo, Brazil KOMAL A. CHANDARANA • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India SAPNA CHANDWANI • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India B. Z. DHOLAKIYA • Department of Chemistry, Sardar Vallabhbhai National Institute of Institute, Surat, Gujarat, India JEAN C. V. DUTRA • Secretaria de Estado da Educac¸a˜o do Espı´rito Santo, Vitoria, Espı´rito Santo, Brazil AMIT GAMIT • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India KEJAL GOHIL • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India MARIA HOLZMANN • Department of Genetics and Evolution, University of Geneva, Geneva, Switzerland G. JAWAHAR • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India SUNIL H. KOLI • Department of Microbiology, Yashwantrao Chavan College of Science, Karad, Maharashtra, India VAIBHAVKUMAR MEHTA • ASPEE SHAKILAM Biotechnology Institute, Navsari Agricultural University, Surat, Gujarat, India BHAVANA V. MOHITE • Department of Microbiology, Bajaj College of Science, Wardha, Maharashtra, India JUN MURASE • Graduate School of Bioagricultural Sciences, Nagoya University, Nagoya, Japan HETVI NAIK • C. G. Bhakta Institute of Biotechnology, Uka Tarsadia University, Surat, Gujarat, India SWATI PATEL • ASPEE SHAKILAM Biotechnology Institute, Navsari Agricultural University, Surat, Gujarat, India ATHARV S. PATIL • H R Patel College of Pharmacy, Shirpur, Maharashtra, India SATISH V. PATIL • School of Life Sciences, Kavayitri Bahinabai Chaudhari North Maharashtra University, Jalgaon, Maharashtra, India VIMAL PRAJAPATI • ASPEE Shakilam Biotechnology Institute, Navsari Agricultural University, Surat, Gujarat, India; ASPEE SHAKILAM Biotechnology Institute, Navsari Agricultural University, Surat, Gujarat, India

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Contributors

VIMALKUMAR PRAJAPATI • ASPEE SHAKILAM Biotechnology Institute, Navsari Agricultural University, Surat, Gujarat, India SHRADDHA SAHA • The Mandvi Education Science College, Veer Narmad South Gujarat University, Surat, Gujarat, India JITENDRA D. SALUNKHE • School of Life Sciences, Kavayitri Bahinabai Chaudhari North Maharashtra University, Jalgaon, Maharashtra, India NARENDRA S. SALUNKHE • School of Life Sciences, Kavayitri Bahinabai Chaudhari North Maharashtra University, Jalgaon, Maharashtra, India MEHREEN SHAH • Department of Environmental Science and Engineering, Jamia Millia Islamia (Central University), New Delhi, India URJITA SHETH • Department of Biotechnology, Faculty of Science, School of Science and Technology, Vanita Vishram Women’s University, Surat, India AMOL D. THENG • Department of Zoology, Bajaj College of Science, Wardha, Maharashtra, India VEYSEL TURAN • Faculty of Agriculture, Department of Soil Science and Plant Nutrition, Bingo¨l University, Bingo¨l, Tu¨rkiye

Part I Abundance and Species Richness Calculation of Protists from Soil

Chapter 1 Soil Sampling, Processing, and Storage Jean C. V. Dutra and Maria C. P. Batitucci Abstract Soil is a natural element that presents unique characteristics, and soil analysis has been the most used technique for diagnosing soil fertility. The organisms that inhabit the soil, such as protists and funghi can be used as bioindicators and can be used to assess environmental changes in a specific area. Soil sampling has been considered a strong tool used to infer about protist populations or communities in the environment. Considering that soil sampling is the basis for the rational, sustainable, and economical use of soils, here we present the key points for carrying out the correct soil sampling, processing, and storage, with emphasis on the sampling process for analysis of soil protists. Key words Soil protists, Soil tracts establishment, Soil sampling, Soil processing, Soil storage

1

Introduction Soil is a natural and three-dimensional element, which occupies a defined section of the landscape and presents unique characteristics. The vertical section of the soil is defined as the soil profile and has its own characteristics that extend from the surface to the material that gave rise to it [1]. On this hand, soil analysis has established itself as the most used technique for diagnosing soil fertility. This technique allows quantifying attributes that benefit and/or harm plant development and stands out for its low operating cost and fast execution [2]. Anthropogenic actions, such as agricultural activity, generate significant impacts on the chemical composition and biota of the soil. In this way, the organisms that inhabit the soil can be used as bioindicators, making it possible to assess environmental changes in a specific area [2, 3]. Numerous biological variables can be quantified in soil samples, such as density, frequency, activity, and biomass of organisms. Thus, analysis of soil biological variables emerges as a strong tool in studies of environmental quality [3].

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Soil biological analyses can be conducted using small amounts of soil maintained in the laboratory or can be conducted in established areas in the research environment. Even under controlled conditions, such as in experiments conducted in the laboratory, collecting and counting all individuals from a population or community in a given area is not feasible. In this way, a sample of the area of interest can be collected, analyzed, and used to infer about a population or community [3, 4]. Activities related to collection, transport, and storage of samples directly influence the results of soil analysis and therefore should not be ignored. Consequently, successful sampling is preceded by careful sampling planning, as this is related to the degree of error in soil analytical procedures. In other words, the representativeness of the samples depends on the sampling design [3]. Considering that soil sampling is the basis for the rational, sustainable, and economical use of soils [1], this chapter will present the key points for carrying out the correct soil sampling, processing, and storage, with emphasis on the sampling process for analysis of soil protists.

2

Materials Contamination is one of the biggest problems of soil sampling. Thus, perform all procedures using clean/sterile materials to avoid contamination. 1. Shovel, 1 L plastic bag, tag for sample identification, sieve [2 mm], 2 L plastic tray. 2. Freezer [-20 °C].

3

Methods

3.1 Soil Tracts Establishment

1. To determine the sampling plots, the site to be studied must undergo a prior analysis of the topography of the land, size and type of particles that make up the soil, type of soil, history of management and vegetation, presence of animals, CO2 and O2 content, moisture content, temperature variation, amount, and distribution of rainfall [5].

3.2 Soil Sample Collection

1. At the sampling site, remove litter, surface vegetation, branches, and stones (do not remove the top layer of soil). 2. With the help of a shovel, collect a composite soil sample from 0–10 cm to 10–20 cm of each tract [6, 7], containing approximately 500 g (see Note 1).

Soil Sampling, Processing, and Storage

5

3. Pack the soil sample in clean plastic bags, tie the opening of the plastic bag together, and attach a label containing the name of the collector and the identification of the sampling tract [3]. 3.3 Soil Processing and Storage

1. Immediately after collecting the composite soil, sieve the soil sample through a 2 mm mesh sieve [6]. 2. Using clean plastic bags, divide the sieved soil into two subsamples. Tie the opening of the bag, and place an identification tag (similar to the one described above). 3. Freeze one of the subsamples at -20 °C, and maintain under these conditions until further soil protist analysis (see Note 2). 4. Air-dry the other subsample in a clean plastic tray, and send for soil chemical analysis (see Notes 3 and 4).

4

Notes 1. Instead of a shovel, an auger can be used to collect the soil sample. 2. It is recommended that the frozen soil sample be analyzed within 4 months. 3. In this step, a part of the sample can be collected to carry out the cultivation of protists present in the soil. 4. Soil chemical analysis recommends soil pH, sum of bases; potential acidity (H + A), cation exchangeability, base saturation, organic matter, macronutrients (phosphorus (P), potassium (K), and calcium (Ca)); and micronutrients (magnesium (Mg), aluminum (Al), iron (Fe), zinc (Zn), cuprum (Cu), manganese (Mn), boron (B), and sulfur (S)). In addition, during the processing of soil samples, the soil exposure to other materials, such as fertilizers, manure and any other elements that may interfere with their composition, must be avoided, since it may interfere in chemical analysis.

Acknowledgments Authors are grateful to UFES (Universidade Federal do Espı´rito Santo) and to FAPES (Fundac¸˜ao de Amparo a` Pesquisa e Inovac¸˜ao do Espı´rito Santo).

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Jean C. V. Dutra and Maria C. P. Batitucci

References 1. Cirilo TF, Machado AOV, da Silva Alves E et al (2020) Diagno´stico do uso do solo na produc¸˜ao agrı´cola no Alto Canal do Serta˜o Alagoano. Braz J Dev 6:52078–52092 2. Brasil EC, Cravo MdS, Veloso AC (2020) Amostragem de solo. Recom adubac¸˜ao e calagem para o Estado Para´ Bele´m, Para´ Embrapa Amaz Orient, pp 47–54 3. Filizola HF, Gomes MAF, de Souza MD (2006) Manual de procedimentos de coleta de amostras em a´reas agrı´colas para ana´lise da qualidade ambiental: solo, a´gua e sedimentos. Embrapa Meio Ambiente, Jaguariu´na

4. Wołejko E, Jabłon´ska-Trypuc´ A, Wydro U et al (2020) Soil biological activity as an indicator of soil pollution with pesticides–a review. Appl Soil Ecol 147:103356 5. Paul EA, Clark FE (1996) Soil microbiology and biochemistry. Academic, San Diego, p 340 6. Oliverio AM, Geisen S, Delgado-Baquerizo M et al (2020) The global-scale distributions of soil protists and their contributions to belowground systems. Sci Adv 6:eaax8787 7. Silva JdS, Fonseca MLdS, Andrade ECd (2022) Metodologia para extrac¸˜ao e detecc¸˜ao de dsRNA em solo

Chapter 2 Initial Observation of Protist from Soil Veysel Turan Abstract Soil is a complex and dynamic ecosystem that is home to a large number of microbial communities. These communities are composed of diverse groups of organisms, including bacteria, fungi, archaea, and protists. Among these, protists are the least studied and recognized organisms in the soil ecosystem, yet they play a significant role in maintaining soil health and ecosystem functioning. Despite their ecological importance, soil protists are often understudied, and relatively little is known about their diversity, distribution, and ecological functions. Recent advances in molecular techniques have helped to shed light on the diversity and role of soil protists in ecosystem processes, and further research in this area may have important implications for the maintenance of soil health and biodiversity. However, the initial observation of soil protists can give a basic idea of the diversity and abundance of the organisms present in the soil. Investigating soil protists is an essential step toward understanding and managing soil ecosystems, and it provides important insights into the complex interactions between microorganisms and soil nutrients. Therefore, the aim of this chapter is to provide an overview of initial observation of soil protists. Generally, two different methods are used to observe naked amoeba and active forms of ciliates and flagellates. Key words Soil protist, Observation, Naked amoeba, Ciliate, Flagellate

1

Introduction Soil protists are single-celled microorganisms found in soil that play a crucial role in the ecosystem [1]. They feed on bacteria [2], fungi [3, 4], and other organic matter [5], regulating nutrient cycling [6], controlling plant pathogens [7], and helping to break down complex compounds and release nutrients [7] back into the soil and that can be accessed by plants [8–10]. Furthermore, soil protists play an important role in maintaining the health and productivity of soils [1, 4]. The relationship of protists with mycorrhizae further improves the plant growth and nutrient content [11]. Overall, soil protists are an essential component and diverse group of unicellular organisms of soil ecosystems and contribute to the functioning and resilience of these systems [12, 13]. Soil protists also help regulate the population sizes of bacteria and fungi, keeping their numbers in

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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check so that they don’t overwhelm the soil [14]. The abundance and diversity of soil protists [15, 16] are important indicators of soil health and microbial community structure, as they play critical roles in nutrient cycling and ecosystem processes [5]. Measuring soil protists can also help in understanding the impact of land-use changes, climate change, or management practices on soil biology and ecological functioning [5, 7, 10, 17]. Soil protists come in a variety of shapes and sizes and are not easily visible to the naked eye [18]. They can be studied using a microscope or DNA sequencing [19]. Soil protists measure refers to the quantification of protists (single-celled eukaryotic organisms) present in soil samples [20, 21]. Some common groups of soil protists include amoebas, flagellates, ciliates, and testate amoebae [6, 17, 22–25]. However, the study of soil protists is challenging due to their small size and diverse morphologies [13, 26]. Nevertheless, advanced microscopy techniques and molecular tools have allowed researchers to better understand the biodiversity and ecological significance of these important microorganisms [1, 4, 18, 20, 21].

2

Materials

2.1 Media and Solutions

1. Non-nutrient agar media: Add 1.5 g agar to 100 mL Page’s saline and autoclave, and pour it in sterile Petri plates (see Note 1). 2. Page’s saline (PAS) (HiMedia). 3. Sterile distilled water.

2.2 Other Requirements

3 3.1

Petri plates, test tubes, inverted microscope, slides, micropipette, or microcapillary.

Methods Soil Sampling

1. Soil samples can be collected from different sites in the desired ecosystem using tools such as trowels, shovels, soil corers, or augers. 2. Collect multiple samples from different locations to get diverse protistan community.

3.2

Common Method

1. Take 1 g of soil sample in a sterile container, and add 10 mL of sterile distilled water in it. 2. Homogenize the sample by vigorous shaking. 3. Allow the soil particles to settle on the bottom of the container.

Initial Observation of Protist from Soil

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4. Using a dropper or pipette, remove a few drops of the liquid from the surface of the container, and place them onto a microscopic slide. 5. Observe the sample under a microscope, using an appropriate magnification (typically 10× or 40×). 6. Focus the microscope and look for moving structures that appear to be protists. 7. Identify the protists present in the sample and take note of their morphology and size. 8. Identify the different types of protists you see and record your observations. 3.3 Observation of Naked Amoeba

1. Follow the steps 1–3 from Subheading 3.2. 2. Take 100 μL soil suspension using a micropipette, and spot inoculate it on previously solidified non-nutrient agar plate. 3. Incubate the Petri plates at room temperature (RT) for 1–2 h (see Note 2). 4. After incubation, observe the Petri plates for slow-moving amoebae under the inverted microscope (see Note 3).

3.4 Observation of Ciliates and Flagellates

1. Weight 5 g of soil sample in Petri plate, and add sufficient water to make a slurry to 80–90% of the soil’s water holding capacity. 2. After 10–15 min, slightly tilt the plate, and firmly press the soil slurry with the thumb. 3. Take out 100 μL (or drop) of the suspension, and place it on the microscopic slide (see Note 4). 4. Observe it under an inverted microscope for active forms of ciliates and flagellates.

4

Notes 1. Make a thin layer of agar which makes observation of species easy. 2. Carry out each step at RT unless specified. 3. The replicate plates can be prepared and incubated at RT in the dark to develop slow-growing amoebae. Observe the plates daily for developing species under an inverted microscope. 4. Pipette out another 100 μL of soil slurry and dilute it up to 1000 μL. From this, add 10 μL of sample to each well of 24 micro-well plate which was previously seeded with 90 uL of PAS solution. Wrap the plates to avoid desiccation and incubate them at RT in the dark for 7 days. Observe the plates under an inverted microscope for presence of small flagellates and active ciliates daily.

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References 1. Geisen S, Mitchell EAD, Adl S, Bonkowski M, Dunthorn M, Ekelund F et al (2018) Soil protists: a fertile frontier in soil biology research. FEMS Microbiol Rev 42:293–323 2. Hawxhurst CJ, Micciulla JL, Bridges CM, Shor M, Gage DJ, Shor LM (2023) Soil protists can actively redistribute beneficial bacteria along medicago truncatula roots. Appl Environ Microbiol 89:e01819–e01822 3. Xiong W, Jousset A, Guo S, Karlsson I, Zhao Q, Wu H, Kowalchuk GA, Shen Q, Li R, Geisen S (2018) Soil protist communities form a dynamic hub in the soil microbiome. ISME 12:634–638 4. Geisen S, Koller R, Hu¨nninghaus M, Dumack K, Urich T, Bonkowski M (2015) The soil food web revisited: diverse and widespread mycophagous soil protists. Soil Biol Biochem 94:10–18 5. Xue P, Minasny B, McBratney A, Jiang Y, Luo Y (2023) Land use effects on soil protists and their top-down regulation on bacteria and fungi in soil profiles. Appl Soil Ecol 185: 104799 6. Dumack K, Feng K, Flues S, Sapp M, Schreiter S, Grosch R, Rose LE, Deng Y, Smalla K, Bonkowski M (2022) What drives the assembly of plant-associated protist microbiomes? Investigating the effects of crop species, soil type and bacterial microbiomes. Protist 173:125913 7. Santos SS, Scho¨ler A, Nielsen TK, Hansen LH, Schloter M, Winding A (2020) Land use as a driver for protist community structure in soils under agricultural use across Europe. Sci Total Environ 717:137228 8. Wang J, Leng P, Shi X, Tan Y, Wang L, Zhang G (2023) Elevated CO2 and/or O3 shift the functional processes and structural complexity of soil protists in a paddy soil. Appl Soil Ecol 185:104806 9. Reehana N, Imran MM, Thajuddin N, Dharumadurai D (2023) Molecular symbiotic interactions of cyanobacterial association in nonvascular seedless plants. In: Microbial symbionts. Academic Press, San Diego, pp 295–309 10. Wang C, Masoudi A, Wang M, Yang J, Yu Z, Liu J (2021) Land-use types shape soil microbial compositions under rapid urbanization in the Xiong’an new area, China. Sci Total Environ 777:145976 11. Henkes GJ, Kandeler E, Marhan S, Scheu S, Bonkowski M (2018) Interactions of mycorrhiza and protists in the rhizosphere

systemically alter microbial community composition, plant shoot-to-root ratio and withinroot system nitrogen allocation. Front Environ Sci 6:117 12. Adl MS, Gupta VS (2006) Protists in soil ecology and forest nutrient cycling. Can J For Res 36:1805–1817 13. Oliverio AM, Geisen S, Delgado-Baquerizo M, Maestre FT, Turner BL, Fierer N (2020) The global-scale distributions of soil protists and their contributions to belowground systems. Sci Adv 6:eaax8787 14. Ren P, Sun A, Jiao X, Shen JP, Yu DT, Li F et al (2023) Predatory protists play predominant roles in suppressing soil-borne fungal pathogens under organic fertilization regimes. Sci Total Environ 863:160986 15. Singer D, Seppey CV, Lentendu G, Dunthorn M, Bass D, Belbahri L et al (2021) Protist taxonomic and functional diversity in soil, freshwater and marine ecosystems. Environ Int 146:106262 16. Malard LA, Mod HK, Guex N, Broennimann O, Yashiro E, Lara E et al (2022) Comparative analysis of diversity and environmental niches of soil bacterial, archaeal, fungal and protist communities reveal niche divergences along environmental gradients in the Alps. Soil Biol Biochem 169:108674 17. Tsyganov AN, Nijs I, Beyens L (2011) Does climate warming stimulate or inhibit soil protist communities? A test on testate amoebae in high-arctic tundra with free-air temperature increase. Protist 162:237–248 18. Geisen S, Bonkowski M (2018) Methodological advances to study the diversity of soil protists and their functioning in soil food webs. Appl Soil Ecol 123:328–333 19. Pellegrino E, Piazza G, Helgason T, Ercoli L (2021) Eukaryotes in soil aggregates across conservation managements: major roles of protists, fungi and taxa linkages in soil structuring and C stock. Soil Biol Biochem 163:108463 20. Geisen S, Mitchell EA, Wilkinson DM, Adl S, Bonkowski M, Brown MW et al (2017) Soil protistology rebooted: 30 fundamental questions to start with. Soil Biol Biochem 111:94– 103 21. Geisen S, Briones MJ, Gan H, Behan-Pelletier VM, Friman VP, de Groot GA et al (2019) A methodological framework to embrace soil biodiversity. Soil Biol Biochem 136:107536 22. Sanders RW (2022) Protists: flagellates and amoebae. In: Encyclopedia of inland waters. Elsevier, London, pp 630–638

Initial Observation of Protist from Soil 23. Venter PC, Nitsche F, Arndt H (2018) The hidden diversity of flagellated protists in soil. Protist 169:432–449 24. Chandarana KA, Pramanik RS, Amaresan N (2022) Interaction between ciliate and plant growth promoting bacteria influences the root structure of rice plants, soil PLFAs and respiration properties. Rhizosphere 21:100466

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25. Pe´rez-Uz B, Galfione VC, Ochoa-Hueso R, Martı´n-Cereceda M (2023) Protist diversity responses to experimental N deposition in biological crusts of a semiarid mediterranean ecosystem. Protist 174:125929 26. Burki F, Sandin MM, Jamy M (2021) Diversity and ecology of protists revealed by metabarcoding. Curr Biol 31:R1267–R1280

Chapter 3 Abundance Calculation of Naked and Testate Amoebae from Soil Urjita Sheth Abstract Protists in soil perform important functions in the decomposer cycle and plant growth and are valued bioindicators for natural and anthropogenic influences. Soil protist includes naked amoebae, testate amoebae, flagellates, and ciliates. Majorly protists in the soil exist in two trophic forms which are vegetative and encysted, that is, the resting stage. Depending upon their stage, the enumeration method for the protist varies. In order to enumerate active forms, direct counting using microscopes is preferred, whereas for encysted protists, various culture methods are reported in the literature. The present chapter has included all the methods used so far for the enumeration and abundance calculation of soil protists. Key words Abundance, Soil protist, Active forms, Encysted protist

1

Introduction Abundance is the measure of determination of biodiversity, which is calculated as the number of individuals found per sample by taking the ratio of the total number of one species to the total number of multiple species present in the ecosystem [1]. Abundance is measured by identifying and counting every individual of each species in a given sector [2]. Protists are defined as single-celled, eukaryotic, heterotrophic, nonfilamentous protists. They are divided into four groups: naked amoebae, ciliates, flagellates, and testate amoebae. Naked amoebae, flagellates, and ciliates feed on bacteria, and these groups are generally abundant in soil. Ciliates are far less numerous in soil than flagellates and amoebae. Flagellates are more abundant in wet soil where relatively large pores are water-filled [3]. In ecological studies, soil protists represent an important group in the soil animal community, and it is always necessary to have information about the numbers of protists with regard to biomass and production [3, 4]. Soil protists have two discrete forms in their

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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life cycle, a trophic that is vegetative and an encysted that is resting stage [5]. Abundance of protists can be determined either by direct method that is without culturing the protist or by indirect method that is by culturing the protist. Determination of active cells without culturing the protist has many merits such as it provides real estimates of active protist, it includes minimum preparative steps, and it is less time-consuming. However, demerits of the method are chances of double counting, chances of misidentification of species, and the samples must be processed within a day or two after collection [6]. The culture techniques [7, 8] are most widely used for the abundance calculations of soil protist [9]. However, some researchers raised questions about the efficiency of culture methods as it determines only active protist [9–11] and recommend direct methods of investigation where a portion of the soil is suspended in water or soil extract and examined under the microscope [4, 11, 12]. The majority of the protists present in the soil are encysted. Using the direct counting method, one can differentiate between active, encysted, and empty testate amoebae easily as their growth rate is slow [4, 13, 14]. Also, testate amoebae can be enumerated precisely by other direct methods too, e.g., the membrane filter technique [15]. The direct counting of ciliates determines the number of active ciliates only [4, 12]. It has often been assumed that direct microscopic examination of soil will yield an unreliable picture of the populations of naked amoebae, heterotrophic flagellates, and ciliates because their numbers are relatively small and they cannot readily be separated from the soil particles [16]. Hence for more abundant small flagellates and amebae, most probable number (MPN) culture methods must be used [16–18]. However, the culture methods do not discriminate among active and encysted forms and only provide the total number of protists in the soil. MPN method is necessary to determine species frequency and biodiversity but cannot be used to determine the abundance of active species in soil [19–21]. This method is not useful for slowgrowing species and those protists that do not feed on bacteria. Also, the growth of protists varies depending upon the duration and condition of storage, sample processing methods, and culturing conditions [4, 22–24]. Because naked amoebae are not readily visible in fresh sediment samples, they were enumerated by enrichment cultivation methods [25] in which a small portion of soil is placed in the Petri dish, and the soil is soaked in water or weak salt solution followed by incubation in the dark at room temperature for few hours and observed under microscope. In this enrichment method, the protist can also be observed directly in the soil-free part of the Petri dish or in a drop of water transferred to a glass slide [3]. However, it is very tough to precisely enumerate heterotrophic flagellates and naked amoebae in soil. It is also tough to determine the proportions of encysted and active individuals [3]. The most

Abundance Calculation of Naked and Testate Amoebae from Soil

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commonly used method for enumerating heterotrophic flagellates and naked amoebae in soil is the so-called most probable number (MPN) modified method of Cutler [7, 8, 17]. This modified method introduced the multi-diluter and the microtiter plate in the counting procedure. Briefly the method includes preparation of dilutions of series of soil suspensions in suitable growth media and observing each dilution at regular time interval for stipulated time period for determining the dilution in which no protistan growth is observed. Using this method, one can estimate the total protist number (active + encysted). The MPN method has certain drawbacks: the method will underestimate total protist numbers if organisms are killed during the setup of the cultures [26] or if they are not able to grow on the food offered. The estimation of the accurate number of protist also varies with the type of media used for the cultivation [27].

2

Materials

2.1 Soil Sample Collection

1. Soil sampling device: scoopula or auger, screw cap plastic tubes and plastic bags, water and alcohol proof marking pen, wash bottle having ethanol, tissue papers. 2. Thermometer, Styrofoam cooler box with ice packs for transport.

2.2 Abundance Calculation of Naked Amoebae

1. 1.5% water agar plates: Add 1.5 g agar to 100 mL distilled water. Mix the contents in an Erlenmeyer flask, melt the agar powder by keeping it in a water bath or microwave, and autoclave for 20 min. Pour the agar in Petri dish to an even thickness of 1.5–2.5 mM, and allow it to cool to solidify (see Note 1). 2. Petri dishes (5 cm diameter), Parafilm, 20 μL or 200 μL pipettor with tips, spatula, aluminum foil. 3. Precision balance, inverted microscope. 4. Fresh soil sample.

2.3 Abundance Calculation of Testate Amoebae

1. Petri dishes (5 cm diameter), distilled water, spatula.

2.4 Abundance Calculation of Testate Amoebae by Direct Count [29, 30]

1. Phenolic aniline blue: Take 15 parts phenol solution (5 g phenol in 100 mL distilled water), 1 part aniline blue solution (dissolve 1 g aniline blue in 100 mL distilled water), and 4 parts concentrated glacial acetic acid. Mix components and filter; stable for years.

2. Precision balance, inverted microscope. 3. Fresh soil sample.

2. Albumen-glycerol.

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Methods

3.1 Collection of Soil Sample

1. For the soil sample collection, first decide your site to be evaluated. 2. Make quadrates of 1 m2 in the proposed field of study. 3. In order to collect the soil sample from the 10 cm depth of the quadrate, with the help of scoopula, take off an entire intact piece of soil plug of 1 cm diameter (diameter which can fit into the sample collection tube) and 10 cm depth (see Note 2). 4. Tag the sample tube aptly. 5. Clean the scoopula by rinsing with ethanol to wash off soil followed by wiping dry with a clean tissue paper. 6. Measure temperature, pH, and moisture content of soil in quadrate at 2 cm and 10 cm depth. 7. In all sampling, collect at least three samples from each quadrate. 8. Put the sampling tubes in a plastic bag and label it properly. 9. Archive one of the three samples in plastic bag in the dark at 4 ° C. 10. From the rest of the two samples, remove above-ground foliage, and using scissors break up and mix the remaining soil. 11. Spread the soil evenly in 15 cm glass Petri dish bases in a containment room at room temperature (18–22 °C) for 6 days. Pass soil through 3.35 mm sieve and homogenize. This material is now referred to as “air-dried soil.” 12. Archive samples in a plastic screw-topped jar. 13. Weigh out 5 g of air-dried soil. Determine its oven-dried weight by heating to 80 °C and weighing at 24 h and 48 h [29]. 14. If one wishes to study the composite sample, collect 6–12 samples by scoopula or cork borer (about 15 mM diameter) from 0 to 3 cm soil depth, in a 4–16 square meter area, and mix the samples thoroughly and withdraw subsamples for the study [28]. 15. Depending upon the nature of the study and the sample size required for the respective study, make other quadrates, and collect samples as described above. 16. Store all the sample bags in the cooler for transportation (see Notes 3 and 4).

Abundance Calculation of Naked and Testate Amoebae from Soil

3.2 Abundance Calculation of Naked Amoebae

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1. Take 1 gram of soil from each of the collected samples in different tubes (see Note 5). 2. Add deionized water to each tube in quantity enough to get the mud-like consistency which can even be pipetted easily. 3. Withdraw 20 μL of the mud and place it onto the agar surface. Put 18–24 drops of mud sample, arranged in rows and columns, onto the agar surface (see Note 6). 4. Label and seal the plates with Parafilm. 5. Incubate the plates overnight in an inverted position (agar side up) in the dark at the temperature of soil at the time of sampling. 6. Repeat the same spotting procedure of the same mud sample using the same pipette tip on a small strip of aluminum foil of known weight. 7. Allow the spots to air-dry. Weigh the foil again and determine the average mean weight of each soil droplet by finding the difference in the weight of foil before placing droplets and after drying droplets. 8. The next day observe plates using the inverted phase-contrast microscope at 200× magnification. Amoebae seem to be at the edge of the soil droplets and begin to migrate across the agar with time. 9. Enumerate the number of amoebae at the edge of each droplet. 10. If the amoebae are in high density, use a square grid in the ocular eyepiece and count representative areas of the edge of the droplet. 11. The samples seen in the plate after the incubation would consist of different encysted amoebae taxa which are reproduced during the overnight incubation due to presence of water. 12. Determine the mean of numbers of the amoebae. 13. Calculate the abundances as “number of cells/gm of dry soil.”

3.3 Abundance Calculation of Testate Amoebae

1. Put 5 mL of distilled or deionized water in a Petri dish. Add 0.5 g of fresh soil to the water, and gently suspend the soil in the plate (see Note 7). 2. Count the number of testate amoebae at the bottom of the plate by observing the plate in an inverted microscope with phase-contrast setting at 50× magnification. Testate amoebae appear at the bottom of the plate in the sediment, and the active species move or explore substrates with pseudopodia (see Notes 8 and 9).

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3. Discard the suspension, clean the plate, and repeat the procedure thrice from the same soil sample collected. 4. Determine the mean of numbers of the amoebae. 5. Calculate the abundances as “number of cells/gm of dry soil.” 3.4 Abundance Calculation of Testate Amoebae by Direct Count [29, 30]

Testate amoebae are usually enumerated by direct microscopy of aqueous soil suspensions [4, 12, 15, 31]. Nowadays modifications of direct counting [13, 32] are used worldwide. 1. Collect composite samples from the site of study (see Note 10). 2. Weigh out 5 g air-dried soil into plastic Petri dish. 3. Add sufficient filtered rain water to produce a slurry. 4. Incubate in the dark at 15 °C for 6 weeks with stirring once in a week. 5. After 6 weeks, take a sample equivalent to 0.5 g of air-dried soil. 6. Place in screw-topped centrifuge tube. 7. Add 7 mL phenolated aniline blue (fixative stain), mix thoroughly, and leave standing overnight (see Note 11). 8. Wash content of storage vessel into a calibrated cylinder, and fill up to 100 mL with distilled water. Close cylinder with Parafilm and mix thoroughly by shaking at least ten times. 9. Take a 1 mL subsample from suspension rapidly after mixing so as to avoid sedimentation. 10. Examine whole subsample by placing suspension dropwise (about 0.1 mL) on grease free slide. 11. Use a compound microscope and a magnification of at least ×100. 12. Full (dark blue stained cytoplasm) and empty tests (unstained or light blue) can easily be distinguished from unstained, inorganic soil particles (see Note 12). 13. Repeat steps 5–8 to record at least 15–30. 14. Calculation: abundances are expressed as number of cells g-1 dry weight of soil. The final calculation of abundance must take into account the initial dilution of soil taken into the Petri dish suspension (e.g.,1 g into 5 mL).

4

Notes 1. Water agar plates should be prepared fresh each time otherwise the agar surface desiccates upon storage. Also, it should be thin enough to ease the visualization of the amoeba in microscope.

Abundance Calculation of Naked and Testate Amoebae from Soil

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2. The collected soil should be added in the tube without forcing or crushing the soil so that the soil porosity, gas exchange, and moisture content can be maintained. 3. Never place the samples directly on ice or in contact with the cold during transportation or in storage for a longer period of time. Try to process the collected samples within a day or 2 from the day of collection, or the species composition might change with the temperature and moisture of the sample which can even lead to change in the bacterial community and eventually overall protistan species composition and relative abundance. 4. If delay in sample processing and samples are to be held in reserve for a long period of weeks or months, air-dry the samples for several days in the refrigerator at 8–10 °C by opening bags and removing the caps of the tubes so as to encyst the protist. Seal the tunes or bags and store at 4 °C. 5. 1 g of the soil sample for determination of naked amoebae should be taken from the core of the soil plug which represents the entire core. 6. The soil sample to be used must be moist so as to prevent desiccation and cell lysis. If the soil sample is sandy, it would become difficult to spot rows of soil. In such a case, soil-water suspension can be spread as a thin layer on the agar surface. 7. Use the amount of solution which can be entirely focused and counted for testate amoebae as in much higher amount of solution, the volume to be scanned would not be focused entirely counts will be difficult to obtain. 8. For too many active individuals in the sample, a 1 cm square grid can be drawn under the plate, or pre-marked grid plates can be used, and one can reduce the amount of soil if the soil texture interferes with observations. 9. To get better estimates of the testate amoebae, one can scan transects through the plate using the inverted phase-contrast microscope at 200× or 400× magnification. For calculations, the number of individuals encountered per transect must be multiplied by the fraction of the plate scanned or the fraction of the soil weight it represents. 10. Usually, 10–20 soil cores are collected from the area studied and thoroughly mixed to a bulk sample. 11. Samples can be stored in this condition for years. Centrifuged tubes with screw tops are ideal for mixing and storing such samples. If suspension becomes colorless after a few hours (sometimes with calcareous soils). Centrifugate sample and replace colorless solution by fresh phenolic aniline blue.

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12. One can add 0.1 mL albumen-glycerol to 1 mL soil suspension if soil particles tend to aggregate on the slide. Preparations should be investigated without coverslip. References 1. Preston FW (1948) The commonness, and rarity, of species. Ecology 29:254–283 2. Verberk WCEP (2011) Explaining general patterns in species abundance and distributions. Nat Educ Knowl 3:38 3. Ekelund F, Rønn R (1994) Notes on protozoa in agricultural soil with emphasis on heterotrophic flagellates and naked amoebae and their ecology. FEMS Microbiol Rev 15:321–353 4. Foisnner W (1987) Soil protozoa: fundamental problems, ecological significance, adaptations in diliates and testaceans, bioindicators and guide to the literature. Prog Protistol 2:69– 212 5. Habte M, Alexander M (1977) Further evidence for the regulation of bacterial populations in soil by protozoa. Arch Microbiol 113: 181–183 6. Carter MR, Gregorich EG (eds) (2007) Soil sampling and methods of analysis. CRC Press, Boca Raton 7. Cutler DW (1920) A method for estimating the number of active protozoa in the soil. J Agric Sci 10:135–143 8. Singh BN (1946) A method of estimating the numbers of soil protozoa, especially amoebae, based on their differential feeding on bacteria. Ann Appl Biol 33:112–119 9. Page AL, Miller RH, Keeney DR (1982) Methods of soil analysis. Part 2. Chemical and microbiological properties, 2nd edn. American Society of Agronomy/Soil Science Society of America, Madison, p 1159 10. Dunger W, Fiedler HJ (1989) Methoden der Bodenbiologie. VEB G 11. Foissner W (1983) Estimation of numbers of protista in soil: a test of the direct method. J Protozool 30:49A 12. Luftenegger G, Petz W, Foissner W, Adam H (1988) The efficiency of a direct counting method in estimating the numbers of microscopic soil organisms. Pedobiologia 31:95–102 13. Couteaux MM (1967) Une technique d’observation des The´camoebiens du sol pour l’estimation de leur densite´ absolue. Rev Ecol Biol Sol 4:593–596 14. Korganova GA, Geltser JG (1977) Stained smears for the study of soil testacida (protozoa, rhizopoda)

15. Lousier JD, Parkinson D (1981) Evaluation of a membrane filter technique to count soil and litter testacea. Soil Biol Biochem 13:209–213 16. Singh BN (1955) Culturing soil protozoa and estimating their numbers in soil. Soil Zool:403–411 17. Darbyshire JF (1974) A rapid micro method for estimating bacterial and protozoan populations. Revue d’Ecologie et de Biologie du Sol 18. Stout JD (1962) An estimation of microfaunal populations in soils and forest litter. J Soil Sci 13:314–320 19. Anderson OR (2000) Abundance of terrestrial gymnamoebae at a northeastern US site: a four-year study, including the El Nino winter of 1997–1998. J Eukaryot Microbiol 47:148– 155 20. Fredslund L, Ekelund F, Jacobsen CS, Johnsen K (2001) Development and application of a Most-probable-number–PCR assay to quantify flagellate populations in soil samples. Appl Environ Microbiol 67:1613–1618 21. Ronn R, Ekelund F, Christensen S (1995) Optimizing soil extract and broth media for MPN-enumeration of naked amoeba and heterotrophic flagellates in soil. Pedobiol 39:10– 19 22. Berthold A, Palzenberger M (1995) Comparison between direct counts of active soil ciliates (protozoa) and most probable number estimates obtained by Singh’s dilution culture method. Biol Fertil Soils 19:348–356 23. Couteaux MM, Palka L (1988) A direct counting method for soil ciliates. Soil Biol Biochem 20:7–10 24. Adl SM (2003) The ecology of soil decomposition. CABI, Wallingford 25. Butler H, Rogerson A (1995) Temporal and spatial abundance of naked amoebae (Gymnamoebae) in marine benthic sediments of the Clyde Sea area, Scotland. J Eukaryot Microbiol 42:724–730 26. Griffiths BS, Ritz K (1988) A technique to extract, enumerate and measure protozoa from mineral soils. Soil Biol Biochem 20:163– 173 27. Rønn R, Ekelund F (1992) The use of different media in estimation of soil protozoan populations by the MPN method. In: Abstract

Abundance Calculation of Naked and Testate Amoebae from Soil No. 108 from the sixth international symposium on microbial ecology, Barcelona 28. Lee JJ, Soldo AT (1992) Protocols in protozoology 29. Finlay BJ, Black HI, Brown S, Clarke KJ, Esteban GF, Hindle RM, Olmo JL, Rollett A, Vickerman K (2000) Estimating the growth potential of the soil protozoan community. Protist 151:69–80 30. Aescht E, Foissner W (1992) Enumerating soil testate amoebae by direct counting. In:

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Protocols in protozoology. Society of Protozoologists/Allen Press, Kansas, pp B6.1–B6.4 31. Meisterfeld R (1989) Die Bedeutung der Protozoen im Kohlenstoffhaushalt eines Kalkbuchenwaldes (Zur Funktion der Fauna in einem ¨ kol 17:221– Mullbuchenwald 3). Verh Ges O 227 32. Couteaux MM (1975) Estimation quantitative des the´camoebiens e´daphiques par rapport a` la surface du sol

Chapter 4 Abundance Calculation of Mycophagous Amoebae Urjita Sheth Abstract Mycophagous protists occur in most soils, but their proper counting methods are lacking. As reported in the literature, mycophagous amoebae can be identified using fungal spores as bait where one needs to culture the mycophagous amoebae with the fungal spores and incubate for several days. In this chapter, two methods are mentioned; one is direct cultivation of amoebae with the spores to find out the lysed and perforated spores, and the other is 24-well microtiter-based method which is known as most probable number (MPN) method. Key words Mycophagous amoebae, Abundance calculation, Culture methods, Microtiter plate technique

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Introduction Obligate mycophagous soil protists include naked amoebae, testate amoebae, ciliates [1, 2], and an obligate, mycophagous soil flagellate [3]. Though mycophagous protist occurs abundantly in most of the soils, their enumeration is not done widely, which might be due to a lack of suitable counting techniques [4]. It is possible to count testate amoebae and actively swimming mycophagous ciliates directly [5]; however, enumeration of mycophagous naked amoebae is conducted by the most probable number (MPN) method on agar plates baited with specific fungi [6, 7]. Here, the use of a particular fungi as protistan feed has certain limitations like the same fungi cannot serve as food for all fungal feeders, and the fungi cultivation is also a tedious procedure [4]. The use of 96and 24-well microtiter plates is reported for the enumeration of mycophagous flagellates [3, 4] using the principle of MPN-method [8] in the presence of Drechmeria coniospora as a food source. However, the use of 96-well microtiter plates can be unreliable for determining the abundance of mycophagous flagellates as the soils usually contain relatively few mycophagous protists, and hence soil particles will mask the protist in the relatively small single wells.

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Therefore, the method proposed here recommends the use of indigenous soil fungi and yeasts in the large wells of 24-well microtiter plates for the determination of mycophagous flagellates.

2

Materials

2.1 Abundance Calculation of Mycophagous Amoebae by Culture Method [9]

1. Soil sample, fungal spores, distilled or deionized water.

2.2 Abundance Calculation of Mycophagous Amoebae Using Microtiter Plates [4]

1. A growth medium for mycophagous protist: Weigh tryptic soy broth (100 mg), glucose (160 mg), modified Neff’s amoebae saline (1000 mL) in an Erlenmeyer flask. Autoclave the mixture, cool it, and add antibiotics to facilitate growth of fungi and yeast but not of bacteria as follows: penicillin (100 mg), streptomycin (150 mg).

2. 250 mL beaker with stir plate and stir bar, 10 mL widemouthed pipette, and 5 mL test tubes. Pasteur pipettes, microscope slides, Petri dishes (35 × 10 mM).

2. Soil sample.

3

Methods

3.1 Abundance Calculation of Mycophagous Amoebae by Culture Method [9]

1. Weigh 10 g of soil sample, and mix it with 90 mL of distilled water in 250 mL beaker using stirrer mixer. 2. Weigh another 10 g of soil sample and air dry at 60 °C for 24 h. This is to determine % soil moisture. 3. Make serial dilutions of the soil suspensions from 101 to 103 times. To do so, withdraw 10 mL of suspension using a large mouth pipette, and add it to another 90 mL of distilled water that is 10-1 dilution. Repeat the same from the 10-1 dilution to get 10-2 and from 10-2 to get 10-3 dilutions. 4. Prepare five aliquots, each containing 1 mL of all of the three dilutions in different test tubes. 5. Allow the soil to settle down. 6. Slowly add 1 mL of fungal spore suspension containing 104 spores/mL to each tube. 7. Allow spores to settle and form a layer on the soil surface. 8. Incubate the tubes for 2–3 weeks at 22–25 °C. 9. After 3 weeks, withdraw sample from each tube consisting of a few drops of spores and soil with a Pasteur pipette, and put the sample on a microscope slide.

Abundance Calculation of Mycophagous Amoebae

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10. Observe using microscope at 250× magnification to check whether the spores are lysed or perforated during the incubation period. 11. Count up to 100 spores/sample and observe for the lysis and perforations. 12. Use MPN techniques and apply a correction factor for soil moisture [10, 11] to estimate the soil population of mycophagous amoeba per gram of dry soil in the original sample (see Note 1). 3.1.1

Observations

1. Observations are made on 100 spores/sample (see Note 2). Normally, 0–3% of spores are lysed because of damage in processing or germination. 2. If 10% or more spores are lysed, the sample is considered positive. 3. Only a few lysed spores with perforations of 5–7 μm in diameter are observed; the sample can be considered positive for Vampyrella, which is the only organism that causes large circular perforations in spores of H. sativum. 4. To confirm the presence of Theratomyxa and Arachnula in the original soil dilution, a subsample containing lysed conidia and soil is added to one side of a Petri dish and baited with fresh spores that are carefully added to the suspension to allow for subsequent observations with a compound microscope at 250×. 5. Digestive cysts containing one and rarely two spores of Helminthosporium indicate the presence of Theratromyxa. 6. Cysts with three to seven spores indicate the presence of Arachnula. 7. Occasionally, more than one species are present in the sample. Therefore, it is important to make observations in a number of locations within the plate. Other species of amoebae may also be observed in these secondary dilution plates as well as interactions among amoeba (see Note 3).

3.2 Abundance Calculation of Mycophagous Amoebae Using Microtiter Plates [4]

1. The soil sample collected should be examined fresh, directly after sampling. 2. Another portion of the same soil should be sieved using 10 × 210) may give negative results. The dilution rate and the number of replications determine the factor for calculating the confidence limits [1].

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Acknowledgment This work was supported by the Japan Society for the Promotion of Science (JSPS) grant 20H02887 to J.M. References 1. Alexander M (1982) Most probable number method for microbial populations. In: Page AL (ed) Methods of soil analysis. American Society of Agronomy, Madison, pp 815–820 2. Singh BN (1946) A method of estimating the numbers of soil protozoa, especially amoebae, based on their differential feeding on bacteria. Ann Appl Biol 33:112–119 3. Murase J, Frenzel P (2008) Selective grazing of methanotrophs by protozoa in a rice field soil. FEMS Microbiol Ecol 65:408–414. https://doi. org/10.1111/j.1574-6941.2008.00511.x 4. Rønn R, Ekelund F, Christensen S (1995) Optimizing soil extract and broth media for MPN-enumeration of naked amoebas and

heterotrophic flagellates in soil. Pedobiologia 39:10–19 5. Page FC (1988) A new key to freshwater and soil gymnamoebae. Freshwater Biological Association, Ambleside 6. Briones AM, Reichardt W (1999) Estimating microbial population counts by ‘most probable number’ using Microsoft ExcelR. J Microbiol Methods 35:157–161 7. Darbyshire JF, Wheatley RE, Greaves MP, Inkson RHE (1974) Rapid micromethod for estimating bacterial and protozoan populations in soil. Revue D Ecologie et de Biologie du Sol 11:465–475

Part III Isolation of Protist from Soil

Chapter 11 Standard Solutions and Media Used for Isolation of Soil Protists Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Abstract Protozoa are heterotrophic, nonfilamentous protists which play a major role in food web decomposition as bacterivores. In both terrestrial and aquatic ecosystems, among all the groups of microbes, they are involved in pivotal processes. Protozoa are conspicuous which may serve as predators and prey in soil ecosystems for other soil microbes. Due to this nature, they may influence the metabolic and development activities of the bacterial communities thereby resulting in increase in plant growth and biomass. Therefore, several researchers have diverted their interest in understanding and identifying the role of protists, and for that, protists need to be isolated and identified. However, for isolation of soil protists, as well as for culturing of isolated protists, several standard solutions and media were used. Key words Isolation, Soil protists, Standard solutions, Media

1

Introduction Protists are essential components of our ecosystem and are considered to be bioindicators of soil quality. Soil protists are of key importance in understanding microbial biogeography and eukaryotic evolution and can be used as potential biofertilizers and biocontrol agents [1]. They consume bacteria, fungi, and other small eukaryotes and therefore occupy a key role in microbial food webs. Predatory soil protists enhance plant growth and production by releasing nutrients into the soil. Protists have an immense morphological and lifestyle diversity [2]. They can be either photoautotrophs such as algae, heterotrophs such as protozoa, or mixotrophs which may obtain carbon phototrophically and heterotrophically [3]. Many protists may form a mutualistic or parasitic symbiotic relationship with plants, animals, fungi, and other protists or may live as host endosymbiotic and/or ectosymbiotic prokaryotes [4]. For isolation, cultivation, and maintenance of protist cultures;

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selective media and liquid solutions were used such as phosphate-buffered soil saline (PBBS), 1.5% agar media, standard soil extract (SSE), 0.1% wheatgrass medium, yeast-beef agar medium (YBA), Prescott-James medium (PJ), non-nutrient agar (NNA), 1% proteose peptone medium, Jone’s medium, and grassseed infusion (GS) [5]. Research in soil biology would be benefitted more by increasing the knowledge about protistology along with the study of bacteria, fungi, and animals [1]. In this book chapter, we presented an overview of the standard solutions and media used for isolation of soil protists so as to use this organism as a model system for understanding the role of these soil protists in improving environmental health and in attaining sustainable agriculture.

2

Material

2.1 Basic Requirements

1. Soil sample for isolating the protists 2. Light microscope/ inverted microscope 3. Glasswares such as Petri dishes, microcapillary, glass pipettes, glass bottles, etc. 4. Antibiotics such as ampicillin, cefoxitin, meropenem, penicillin, and streptomycin

2.2

Media

1. Phosphate-buffered saline (PBS) (1X, pH 7.4): First of all, pour 800 mL of distilled water in a suitable container such as glass flask. Weigh 8 g of 137 mM NaCl, 0.2 g of 2.7 mM KCl, 1.44 g of 10 mM Na2HPO4, 0.24 g of 2 mM KH2PO4, 0.0476 g of 0.5 mM MgCl2, and 0.147 g of 1 mM CaCl2. Now, add all these weighed salts to 800 mL distilled water in a flask with continuous stirring. Adjust pH of the soil with 1 M HCl from which the cells were obtained. Top up with distilled water to make it 1 L. Keep the cell suspensions in this buffer for a few hours. Use this solution to hold cells in suspension or for washing cells. Instead of using distilled water or deionized water, cells get less damaged using this solution [5]. 2. 1.5% Agar: Pour 100 mL of distilled or deionized water in an Erlenmeyer flask, and add 1.5 g of agar powder to it. Autoclave the media for 20 min in an autoclave. Prepare the agar by substituting the water with standard soil solution (SSS) or standard soil extract (SSE) or wheatgrass medium [5]. 3. Standard soil extract (SSE): This solution contains dissolved nutrients from the soil sample. This soil extract is used as a supplement for any of the growth media but cannot be used as a growth medium itself (see Note 1). To prepare SSE, add 300 g of soil from A horizon and 1 mL distilled water or

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deionized water in a large beaker, and stir continuously for 1 h. Allow the soil to settle for 30 min, and then filter the soil extract through several layers of cheesecloth in an Erlenmeyer flask. Dispense the filtered extract into screw cap tubes, and then autoclave them for 20 min. 4. Standard soil solution (SSS): SSS is a solution where concentration of ions is known. Weigh 6.8 × 10-4 g/L KH2PO4, 1.116 g/L FeCl3, 0.241 g/L MgSO4, 0.544 g/L CaSO4, 0.133 g/L NH4Cl, 0.253 g/L KNO3, and 0.14 g/L NaCl salts, and dissolve these salts one at a time in 900 mL distilled water with continuous stirring. pH of solution should be adjusted to desired level using phosphate buffer, and the pH should be pH ± 0.3 as of the soil from which the cells are obtained. After adjusting the pH of the solution, add distilled water to make up the final volume to 1 L, and then dispense the solution into screw cap tubes. Sterilize the solution by autoclaving it for 20 min, and then store the solution at 4 °C. 5. 0.1% wheatgrass medium: This media can be used to culture many protozoan species. Take 1 g wheatgrass powder, dissolve it in 1 L distilled or deionized water, and boil it to allow to infuse at a gentle rolling boil for 2 min. Let the media settle down and allow to cool for an hour. Filter the media into a new flask by using several layers of cheesecloth. Discard the grass residue, and adjust the pH (pH ± 0.3) to that of the soil under study using phosphate buffer. Autoclave the media for 20 min, and dispense the media into screw cap tubes. This medium can be prepared with SSS or SSE instead of deionized or distilled water. To reduce bacterial growth, this media can be diluted to 1/10 or 1/100 times, and many protozoan species can be observed when bacterial growth is low. 6. Yeast-beef agar medium (YBA): Dissolve 0.5 g yeast extract powder, 0.5 g beef extract powder, and 1.5 g agar powder in 100 mL distilled or deionized water in an Erlenmeyer flask. Autoclave the media for 15 min (see Note 2) [5]. 7. Non-nutrient agar (NNA): Prepare this media by adding 1 L of Neff’s amoeba saline (AS) and 15 g of non-nutrient agar. Boil it. 8. Protein-rich liquid media: Protein-rich liquid media are mostly preferred for isolation and cultivation of protists axenically and for some cultures, Jones’s medium containing bacteria is preferred. 9. Proteose peptone glucose (PPG): Dissolve 10 g proteose peptone and 18 g glucose in 1 L of Neff’s amoeba saline (AS) and then sterilize the media in culture tubes by autoclaving it.

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10. Chang’s serum/casein/glucose/yeast extract medium (SCGYEM): Take 10 g of isoelectric casein, 2.5 g glucose, 5 g yeast extract, 1.325 g Na2HPO4, 0.8 g KH2PO4, and 100 mL fetal calf serum, and dissolve all these ingredients one at a time in 900 mL distilled or deionized water. If the inoculum contains bacteria, add 200 μg/mL of antibiotic such as penicillin and streptomycin. Sterilize the media by autoclaving the media in culture tubes. 11. Jones’s medium: Dissolve 2.65 g Na2HPO4.12H2O, 0.41 g KH2PO4, and 7.36 g NaCl in 1 L of distilled or deionized water, and adjust the pH to 7.2. The media should contain 850 mL buffered saline, 50 mL horse serum, and 100 mL of 1% yeast extract solution. Autoclave the media, pour it into the sterile tubes, and add a pinch of sterile rice starch to each tube (see Note 3) [6].

3

Methods 1. Collect the soil samples from natural habitats from which the protists are needed to be isolated with a sterile equipment in order to avoid any contamination (see Note 4). 2. Immediately, after transporting soil samples to the laboratory, examine the soil samples using an inverted microscope for visualization of the protist species. 3. Inoculate the soil samples to the selective medium such as phosphate-buffered soil saline (PBBS), 1.5% agar media, standard soil extract (SSE), 0.1% wheatgrass medium, yeast-beef agar medium (YBA), Prescott-James medium (PJ), non-nutrient agar (NNA), 1% proteose peptone medium, and grass-seed infusion (GS) according to the protozoan species which are sought to be isolated, for example, proteose peptone medium to isolate Tetrahymena spp.; wheatgrass medium to grow protozoa such as rhizopods, choanoflagellates, ciliates, and flagellates; Prescott-James medium (PJ); non-nutrient agar (NNA); grass-seed infusion for naked amoebae; and 1.5% agar medium and soil extract medium for isolating Acanthamoeba spp. 4. For isolating the protozoan species, incubate the cultures at 37 °C for 7–8 days (see Note 5). 5. Subculture the samples into the appropriate medium for enrichment of the protozoan species, and for subculturing, viable cell count should be numerous.

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6. Along with the protozoa, bacteria or small protists which may serve as food for the protozoa like large amoebae need to be eliminated if one has to cultivate the protists axenically to study the biochemical, genetic, physiological, molecular, and taxonomic studies. 7. Inoculate the antibiotics such as ampicillin, cefoxitin, meropenem, penicillin, and streptomycin into the media to subculture the protozoan species axenically [6].

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Notes 1. With each preparation, composition of the solution varies according to the soil used. It is advisable to provide a solution with ions similar to the soil used. 2. Avoid excessive sterilization so as to prevent the accumulation of toxic denatured protein derivatives. Try to use the lowest autoclave temperature to ensure sterility. 3. Heat the rice starch at 150 °C for 2 h to ensure sterility. 4. Samples should be collected with sterile equipment before inoculating the samples into the appropriate media. 5. Incubate the plates in the normal position to ensure that the suspension does not run down onto the lid of the plate.

References 1. Geisen S, Mitchell EA, Adl S, Bonkowski M, Dunthorn M, Ekelund F et al (2018) Soil protists: a fertile frontier in soil biology research. FEMS Microbiol Rev 42(3):293–323 2. Caron DA, Worden AZ, Countway PD, Demir E, Heidelberg KB (2009) Protists are microbes too: a perspective. ISME J 3:4–12 3. Geisen S, Bonkowski M (2018) Methodological advances to study the diversity of soil protists and their functioning in soil food webs. Appl Soil Ecol 123:328–333

4. De Vargas C, Audic S, Henry N, Decelle J, Mahe´ F, Logares R, Lara E et al (2015) Eukaryotic plankton diversity in the sunlit ocean. Science 348:1261605 5. Carter MR, Gregorich EG (eds) (2007) Soil sampling and methods of analysis, 2nd edn. CRC Press, New York 6. Kalinina LV, Page FC (1992) Culture and preservation of naked amoebae. Acta Protozool 31: 115–126

Chapter 12 Isolation Techniques for Amoeba Jun Murase Abstract Amoebae are among the most common and abundant protists in all soil types. They can be relatively easily isolated using simple methods when they are cultivable. Here, I describe three methods commonly used for the isolation of phagotrophic amoeba: a migration method, pipette technique, and dilution technique. Key words Cultivation, Dilution method, Migration, Pipetting, Soil amoeba

1

Introduction Identification of an amoeba species requires the establishment of a clone. After recognizing an amoeba morphotype in an initial culture, the amoeba should be cloned before further investigation to avoid errors that would arise if the culture consisted of a mixture of similar amoeba species [1]. Various methods for cloning amoebae have been suggested [2], among which a migration method, pipette technique, and dilution technique are the three primary methods [1]. A migration method is applicable for amoebae that grow on agar media, and it is relatively easy to obtain a clone. A pipette technique is a method for cultures growing in liquid media and requires practice. A dilution technique is simple but based on the prerequisite that one of the dilutions should contain only one or very few cells.

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Materials

2.1 Amoeba Saline Solution (Modified Neff’s Amoeba Saline, [2]) (See Note 1)

1. Prepare 100× stock solutions for each element: 1.20 g NaCl, 0.04 g MgSO4·7H2O, 0.04 g CaCl2·2H2O, 3.58 g NaHPO4·12H2O, and 1.36 g KH2·PO4 per 100 mL.

2.2 1% Agar Amoeba Saline Media

1. Add 1 g agar to 100 mL amoeba saline and autoclave it.

2.3 Food Bacteria Suspension

1. Culture Escherichia coli in sterilized liquid organic media like nutrient broth at 37 °C overnight (see Note 3).

2. Add 10 mL of each stock solution to 950 mL of water and autoclave it.

2. Pour 1 mL of melted agar into a tissue culture dish (diameter 35 mm) to make a thin agar layer (see Note 2).

2. Harvest them by repeated centrifugation (10,000 g, 10 min, 4 °C) and washing with sterilized amoeba saline. 3. Resuspend the bacterial cells in amoeba saline solution. Determine the cell density under the microscope using a Thoma cell counting chamber. This stock can be stored for a month in the refrigerator. 4. Dilute the suspension to 108 cells mL-1 using sterile amoeba saline before use. 2.4 96-Well Microplate

A flat-bottom type is better for observing the growth of amoeba under an inverted microscope (e.g., CAT No. 167008, Thermo Fisher Scientific).

2.5 Fine Capillary Pipette

After heating in a flame, pull the end of a Pasteur pipette into a fine capillary tube.

2.6

A small scalpel is appropriate for cutting out a small agar media (see Note 4).

3

Scalpel

Methods Carry out all procedures at room temperature unless otherwise specified.

3.1 Preparation of a Soil Dilution Series

1. Mix 10 g of a soil sample, collected using sterile instruments, with 90 mL of amoeba saline. 2. Shake the suspension using a rotator for 10 min. 3. Mix the soil suspension with amoeba saline at a 1:1 ratio, and continue to prepare a threefold dilution series: from 10 × 20 up to 10 × 211 (see Note 5).

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Incubation

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1. Distribute the diluted suspensions (50 μL, eight replicates) to a 96-well microplate, and add 100 μL of food bacteria suspension. 2. Incubate the plates at a temperature close to the in situ soil temperature in the dark.

3.3 Subculturing Amoeba

1. Observe the growth of phagotrophic amoeba in the wells under an inverted microscope at 200× magnification using phase contrast optics. 2. Transfer trophozoites/cysts in the wells to a new food bacteria suspension in a 96-well microplate or tissue culture plate, and incubate them. 3. Repeat steps 1 and 2 several times.

3.4 Isolation of Amoeba 3.4.1

Migration Method

1. Distribute 1 mL of food bacteria suspension on 1% agar amoeba saline media. After settling down bacteria on agar for 1 h, remove the overlying liquid. Keep the media in an incubator overnight to dry the surface. 2. Spot an aliquot of amoeba subculture obtained in Subheading 3.3 (ca. 10 uL) on the edge of the media (see Note 6). 3. Incubate the spotted amoeba subculture. 4. Observe the media under an inverted microscope, and find areas where amoeba have migrated away from the initial path of the inoculation and are not too abundant. 5. Cut out one amoeba or cyst on a small block of agar with a scalpel. 6. Transfer the block to the center of a fresh agar media, and wash the cell with a drop of amoeba saline. 7. Incubate the subculture and observe the growth and morphotype of amoeba. 8. Repeat steps 3–6 until a single morphotype of amoeba cells is obtained.

3.4.2

Pipette Technique

1. Observe the subculture obtained in Subheading 3.3 under an inverted microscope. 2. Transfer one amoeba or cyst to a fresh food bacteria suspension using a fine capillary pipette under a microscope. 3. Incubate it, and repeat steps 1 and 2 when a single morphotype of amoeba cells is obtained.

3.4.3

Dilution Technique

1. Wash the amoeba in subculture in Subheading 3.3 using fresh amoeba saline. 2. Dilute the subculture at appropriate rates (see Note 7).

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3. Transfer the aliquots of dilutions to a new food bacteria suspension. 4. Incubate the subculture and observe the growth and morphotype of amoeba. 5. Repeat steps 2–4 until a single morphotype of amoeba cells is obtained.

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Notes 1. Other saline solutions (Page 1988) can also be used. 2. A thin agar layer enables observation of protists growing on agar under an inverted microscope. 3. Other Enterobacteriaceae bacteria, such as Enterobacter and Klebsiella, can be used as alternative food bacteria. 4. A micro spatula with a flat end can also be used. 5. The degree of dilution can be modified; narrower dilution ranges or a fixed dilution rate can be adapted when the estimated number of amoebae in the soil is available. 6. Provide sufficient space for the spotted amoeba to migrate over the media. 7. Enumeration of the cell number using a cell counting chamber (e.g., improved Neubauer’s counting chamber) helps determine the dilution rates.

Acknowledgment This work was supported by the Japan Society for the Promotion of Science (JSPS) grant 20H02887 to J.M. References 1. Smirnov AV, Brown S (2004) Guide to the methods of study and identification of soil gymnamoebae. Protistology 3:148–190 2. Page FC (1988) A new key to freshwater and soil gymnamoebae. Freshwater Biological Association, Ambleside

Chapter 13 Isolation of Ciliates and Flagellates from Soil Amol D. Theng, Bhavana V. Mohite, and Satish V. Patil Abstract Protists are unicellular eukaryotes consisting of various forms like ciliates, flagellates, and ameboid. Ciliates are one of the distinct groups of protozoans characterized by cilia for locomotion, sensing, as well as feeding, while flagellates are unicellular organisms having whip-like lashing structures called flagella. Ciliates are structurally very simple which makes them highly sensitive to environmental changes which make them useful bioindicators of any ecosystem. Flagellates play an important role in the aquatic food chain by its role in nutrient cycling as primary producers as well as consumers of bacteria, algae, and other microorganisms. Ciliates are relatively difficult to isolate and to culture in laboratory conditions. Generally, protists can be retrieved from the soil by the direct or indirect method. The present protocol describes thorough methodology for isolation and enumeration of ciliates and flagellates with its microscopic observation and brief about identification. Key words Excystment, Bioindicator, Food web, Heterotrophs

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Introduction Protists are unicellular eukaryotes. Heterotrophic forms of protists consist of various forms like ciliates, flagellates, and ameboid. All these forms are essential components of aquatic and soil ecosystem [1]. Ciliates Ciliates are one of the distinct groups of protozoans characterized by cilia for locomotion, sensing as well as feeding. ciliate size ranges from 10 μm to 4 mm in size. Nearly 3000 species of free-living ciliates are estimated [2], while Liu et al. [3] estimated underperformed identification of global ciliates which is around 30,000 species. Unlike other eukaryotes ciliates are dikaryotic, i.e., they carry two nuclei a large, polyploid macronucleus or vegetative nucleus and a tiny, diploid micronucleus or generative nucleus.

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Ciliates are found in diversified habitats ranging from freshwater, marine water, soil, heavy metal contaminated sites, deep sea, and puddles showing their cosmopolitan distribution. Most of the ciliates form cysts; a dormant phase that can remain viable for many years under unfavorable conditions like desiccation and frost [4]. They remain viable until the environmental conditions become favorable for excystment. Ciliates have higher growth rates than others often marking them as a contaminant in the algal system [5]. Ciliates are important links in the aquatic food chain and are accountable for consuming 10–80% of phytoplankton’s primary production in marine ecosystems [6]. Ciliates reproduce asexually by fission and sexually by conjugation. Asexual reproduction is the main mechanism through which ciliates increase their population size during their life cycle [7]. Protists, especially ciliates, are structurally very simple which makes them highly sensitive to environmental changes as compared to other forms like amoebae [8]. On the other hand, they are present at the primary consumer level in the aquatic food chain; these facts make them useful bioindicators of any ecosystem [9]. Ciliates can also be used in bioassays to monitor heavy metal pollution [10]. Ciliates are relatively difficult to isolate and to culture in laboratory conditions [11]. Generally, protists can be retrieved from the soil by the direct or indirect method. The direct method works out well when moist and freshly drawn soil samples are used. On the other hand, indirect methods allow the activation of inactive cysts. Flagellates Flagellates are unicellular organisms having whip-like lashing structures called flagella; they may range from one to many depending on the species. Flagella are used for movement and sensory purposes as well as directing food for ingestion [12]. They are widely distributed in aquatic habitats, like freshwater, marine, and brackish water. Most of them are free-living and some are parasites. These organisms are mostly photosynthetic, but some are heterotrophic. They play an important role in the aquatic food chain by being primary producers as well as consumers of bacteria, algae, and other microorganisms. Flagellates also play a crucial role in nutrient cycling (see Note 1). Heterotrophic flagellates feed on bacteria and dissolved organic matter [13]. They are found widespread in soils with great abundance [14]. Functionally, they link the carbon and nutrient flow between primary producers with higher trophic levels, making them very important players in the microbial food web in soil [15].

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Materials 1. Soil sample, sugar, yeast, double distilled water 2. Dropper, Petri plates, beaker, glass container, glass slides, coverslips, spatula, micropipette with disposable tips 3. Light/phase contrast microscope, precision balance

3

Methods

3.1 Collection of Soil Sample

1. Collect 50 g of soil sample in a clean plastic container from the soil preferably beneath 5–10 cm from soil surface. Spatula used for soil collection should be sterilized to avoid crosscontamination. 2. From finely powdered or stone-free sample, 10 g of soil should be utilized for further processing. 3. Along with sample collection mark parameters like temperature, moisture, and pH of soil. 4. Sample must be registered for collection time, date, and site coordinates. Sample should be manually screened for any other physical contaminants. 5. Moist/wet samples should be processed within 24 h of collection, while dry soil samples should be kept in an airtight container till processing. (see Notes 2–4)

3.2 Incubation of Soil Sample

1. Add 5 g of soil sample to 250 mL of distilled water (autoclaved, so as to minimize waterborne entry of protists). 2. Sample bottles should be closed and kept at room temperature (30 °C) in a partially illuminated area avoiding direct sunlight for 24 h. 3. After 48 h, take a drop of sample from jar on a glass slide with a coverslip, and observe under the microscope. 4. The culture should be observed every few days for species activity. A suggested observation schedule is on 2, 4, 6, 10, 14, and 20 days [16]. 5. On the other hand, estimation of active cells that employ the most probable number (MPN) approach involves the preparation of replicating dilution series in a culture medium and counting on successive days [17].

3.3 Enumeration of Ciliates and Flagellates

1. Active ciliates appeared quickly as soon as the collection of soil samples, but activation of cysts requires time from at least 4–10 days. Active ciliate from wet soil can be observed immediately after the collection of soil (see Note 2).

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2. Pipette 15 μL of suspension from incubated stock soil solution to the hemocytometer chamber, and cover. 3. Scan the counting chamber with 200× and 400× under phase contrast 4. Repeat the procedure 3–5 times until a stable mean abundance value is obtained. 5. If initial stock soil solution has high abundance or soil particles which make enumeration difficult, serial dilution of stock solution is recommended (see Note 5). 6. The final calculation of abundance must take into account the initial dilution of soil into the Petri dish suspension. 3.4

Calculations

Calculations for moist and dry soil can be made as per the method cited by Carter et al. [16].

Abundance of ciliates=flagellates per g dry soil = ðAbundance in each filter × conversion factorÞ=dry weight soil in Petri dish where, Abundance in each filter = Total no. of ciliates/flagellates measured in a Hemocytometer Conversion factor = dilution factor or fraction of sample used out of total stock solution Dry weight of soil in the Petri dish = ðwet weight ðgÞ × 100Þ=100 þθ where θ is % of moisture content of soil. 3.5 Fixing and Staining

Staining of protists is equally important to differentiate minute differences, and a variety of stains can be used. 1. Methylene blue (0.1% (w/v) in distilled water) will color the nuclei dark blue against a light blue cytoplasm. 2. Neutral red (0.1% (w/v) in distilled water) is a cytoplasmic stain. 3. Sudan III and Sudan Black stains lipid droplets. 4. 4′-6-Diamidinob-2-phenylindole (DAPI) is a nuclear stain (see Note 6). 5. Klein’s “Dry” Silver will stain cilia, flagella, and cell surface [18]. 6. Protargol is a silver proteinate stain, which stains cilia as well as internal structures such as microtubuli, extrusomes, and nuclei [19] ( Fig. 1).

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Fig. 1 Representative example of ciliates and flagellates under 40× light microscope (a) Euplotes sp. (b) Vorticella sp. (c) Paramecium sp. 3.6

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Identification

Identification of protists is one of the toughest tasks due to morphological resemblance. Microscopic observation, staining, as well as molecular approaches can be used. During the counting process, the species diversity of ciliates was determined by identifying observed ciliates using taxonomic keys [20, 21] (see Notes 7 and 8).

Notes 1. Flagellates are very abundant in soils. Most are bacterivores showing predation preference for specific bacterial species. Thus, for culturing flagellates one should be very cautious about the availability of specific food. 2. For direct counts of active species, samples must be processed within a day or two. Species composition will change as temperature or moisture in the sample changes. Typically, bacteria and therefore the bacterivore community will change, and this will also affect overall protozoan species composition and relative abundances. 3. If samples are to be stored for a long time (weeks or months), it is recommended to let the soil air dry slowly over several days. When sufficiently dry, containers can be sealed and transferred to 4 °C. 4. As the only reliable method for estimating in situ abundance of active (i.e., non-cyst) protozoa, the number of active cells can only be determined without culturing. In most cases, dry soils do not report any active organisms, leading to low taxonomic resolution.

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5. Typically, there are 105–107 flagellates per gram of dry soil. In this scenario with such abundance, it’s difficult to count accurately with a counting chamber. It is recommended to use most probable number of enumerations with keeping record of serial dilutions. 6. The stock solution of 4′-6-diamidinob-2-phenylindole (DAPI) stain is made by preparing a 10 μg μL-1 solution in distilled water. This stock is stored at -20 °C. The stock is diluted 1: 1500 or 1:2000 before staining. The usual final stain concentration is 0.5 μg mL-1. 7. In addition to microscopic observation, molecular techniques can be used to identify ciliates. Due to the high morphological similarity between species, identifying ciliates based on microscopic observations of living and/or preserved individuals is often challenging [11]. 8. Identification of protists needs to be confirmed by molecular sequencing by 18s rRNA sequencing analysis.

Acknowledgment Author ADT and BVM are grateful to Principal, Bajaj College of Science, Wardha, for the encouragement and support. References 1. Adl S, Coleman D (2005) Dynamics of soil protozoa using a direct count method. Biol Fertil 42:168–171 2. Finlay BJ, Esteban GF (1998) Freshwater protozoa: biodiversity and ecological function. Biodivers Conserv 7:1163–1186 3. Liu W, Fan X, Jung JH, Grattepanche JD (2022) Editorial: ciliates: key organisms in aquatic environments. Front Microbiol 13: 880871 4. Foissner W, Agatha S, Berger H (2002) Soil ciliates (Protozoa Ciliophora) from Namibia (Southwest Africa), with emphasis on two contrasting environments, the Etosha Region and Namib desert. Denisia 5:1–1459 5. Smith VH, Crews T (2014) Applying ecological principles of crop cultivation in large-scale algal biomass production. Algal Res 4:23–34 6. Rassoulzadegan M, Cowie A, Carr A, Glaichenhaus N, Kamen R, Cuzin F (1982) The roles of individual polyoma virus early proteins in oncogenic transformation. Nature 300:713–718

7. Lynn DH (ed) (2008) The ciliated protozoa: characterization, classification, and guide to the literature. Springer, New York 8. Petz W, Foissner W (1989) The effects of mancozeb and lindane on the soil microfauna of a spruce forest: a field study using a completely randomized block design. Biol Fertile Soil 7: 225–231 9. Payne RJ (2013) Seven reasons why protists make useful bioindicators. Acta Protozool 52: 105–113 10. Gutie´rrez JC, Amaro F, Martı´n-Gonza´lez A (2009) From heavy metal-binders to biosensors: ciliate metallothioneins discussed. Bioessays 31:805–816 11. Duff RJ, Ball H, Lavrentyev PJ (2008) Application of combined morphological–molecular approaches to the identification of planktonic protists from environmental samples. J Eukaryot Microbiol 55:306–312 12. Mitchell DR (2007) The evolution of eukaryotic cilia and flagella as motile and sensory organelles. In: Eukaryotic membranes and

Isolation of Ciliates and Flagellates from Soil cytoskeleton, origins and evolution. Springer, New York, pp 130–140 13. Leadbeater BS, Green JC (2000) Flagellates: unity, diversity and evolution. CRC Press, Boca Raton 14. Geisen S, Tveit AT, Clark IM, Richter A, Svenning MM, Bonkowski M, Urich T (2015) Metatranscriptomic census of active protists in soils. ISME J 9:2178–2190 15. Geisen S, Bonkowski M (2018) Methodological advances to study the diversity of soil protists and their functioning in soil food webs. Appl Soil Ecol 123:328–333 16. Carter M, Gregorich E, Adl S, AcostaMercado D, Anderson T, Lynn D (2007) Methods in soil protozoa. In: Soil sampling and methods of analysis. CRC Press, Boca Raton

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17. Fredslund L, Ekelund F, Jacobsen CS, Johnsen K (2001) Development and application of a Most-Probable-Number–PCR assay to quantify flagellate populations in soil samples. Appl Environ Microbiol 67(4):1613–1618 18. Fariya N (2018) Protocol identification and preservation of myxozoan parasites for microscopy with silver nitrate (Klein’s dry) staining technique. Microsc Res Tech 81:1162–1164 19. Lynn DH (1992) Protargol staining. Protocols in protozoology. Soc Protozool Lawrence C 4: 1–C4 20. Foissner W (1998) An updated compilation of world soil ciliates (Protozoa, Ciliophora), with ecological notes, new records, and descriptions of new species. Eur J Protistol 34:195–235 21. Foissner W (2016) Protists as bioindicators in activated sludge: identification, ecology and future needs. Eur J Protistol 55:75–94

Chapter 14 Isolation and Enumeration of Mycophagous Protist Amol D. Theng, Bhavana V. Mohite, and Satish V. Patil Abstract Protists can prey upon other smaller organisms such as bacteria and fungi. Their feeding actions are of utmost importance in driving the microbial loop and food web. The bacterial communities are suggested as their main prey, but it is not the sole major determinant. Mycophagous/fungivore protists are more widespread and abundant in different soils and could be ciliates, flagellates, and amoeba. Mycophagy is widespread due to development of specialized enzyme system to digest fungi. The mycophagous protists developed a specialized structure like a feeding tube to perforate the cell wall of yeasts, fungal hyphae, and spores to take up their content. Diverse morphologies of yeast and filamentous fungi induce distinct killing and feeding mechanisms in mycophagous protists. Mycophagous protists constitute a reservoir of biocontrol agents that could directly consume fungal pathogens. The present protocol summarizes the isolation and enumeration of mycophagous protists. Key words Soil biodiversity, Food web, Mycophagy, Fungal energy channel

1

Introduction When the role of protists within the rhizosphere microbiome was studied, protists function as “puppet masters,” the beneficial plant– microbe interactions exploited to operate the rhizosphere microbiome functionality [1]. Soil protists play an important role in the food chain, and their diversity is a general marker of soil quality. On the basis of the mode of nutrition, soil protists are autotrophs or heterotrophs. Autotrophic soil protists are photoautotrophs that produce their own food from inorganic substances, like nitrates and ammonia. Heterotrophic soil protists can only use organic substances to produce food. They are also called chemoautotrophs. Heterotrophs can be broadly utilizing two modes of nutrition either bacterial or fungal energy channels. Fungal Energy Channel Soil biologists generally classify the nutrient flow in soil food webs into a bacterial and a fungal-based energy channel. Fungal energy

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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flow often highlights arthropods and nematodes, but there are many studies revealing an important share of mycophagous protists. There were many facultative and obligate mycophagous protists found in soil [2, 3]. Mycophagy is reported in ciliates [4], flagellates [5], and amoeba [6]. Although bacterivorous protists are considered as major consumers at tropic level, mycophagous protists can reach biomasses similar to them [3]. The major reason behind the underrated importance of mycophagous protists might be due to the unavailability of specific isolation and cultivation methods and trending methods of isolation that select for bacterivorous protists [7, 8]. Mycophagous Protist and Soil Protist taxonomists have long realized that diverse facultative and obligate mycophagous protist taxa are common in soils [2, 3, 9]. Recent studies found that mycophagous are more widespread and abundant in different soils than previously taught [10]. Widespread distribution of mycophagous protists suggests presence of specialized enzyme system to digest fungi while producing secondary metabolites that prevent growth of bacteria and thus prevent resource partitioning. On the contrary, soil bacteria are also evolving to cope with those toxic secondary metabolites and acquire specific energy resources by competing [11, 12]. These facts make bacteria and fungi co-occur in same habitat and thus bacteriophagy and mycophagy seen commonly in soil protists [13]. The mycophagous protists like ciliates grossglockneriids developed a feeding tube to perforate the cell wall of yeasts, fungal hyphae, and spores to take up their contents [14]. The different morphologies of yeast and filamentous fungi trigger distinct killing and feeding mechanisms in a fungivorous amoeba [15]. Moreover, several species previously considered bacterivorous were recently discovered to feed on a range of fungal spores and yeast cells, including plant pathogenic fungi [16]. This widespread mycophagy suggests that mycophagous protists constitute a reservoir of biocontrol agents that could directly consume fungal pathogens. Protists now look as microbiome optimization tool for sustainable agriculture [1] (Fig. 1).

2

Material 1. Soil sample 2. Yeast culture (e.g., Saccharomyces sp., Candida sp.)/fungal spore (e.g., Aspergillus sp.) 3. Micropipette, glass bottles, 24-well microtiter plate 4. Phase contrast microscope

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Fig. 1 Mycophagous protists: types, mechanism, and significance

5. Preparation of growth medium: Add 100 mg tryptic soy broth and 160 mg glucose in 1000 mL modified Neff’s amoeba saline, adjust pH 6.8–7.9 [7]. After autoclaving and cooling, add 100 mg penicillin and 150 mg streptomycin (see Notes 1 and 2).

3

Methods

3.1 Isolation of Mycophagous Protists

Isolation of mycophagous protists can be carried out by following the method as described by Ekelund et al. [3] with slight modifications. 1. Take 5.0 g of soil sample and suspend it in 100 mL growth medium in a glass bottle. 2. Shake these bottles on shaker for 30 min. 3. After shaking, add 2 mL of the suspension to a test tube containing 6 mL fungal growth medium. This results in fourfold dilution. 4. Vortex the solution for 10 s. 5. Repeat above step (steps 3 and 4) five times, so, a total of six dilutions will be there.

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6. From each set of the six dilutions, take 1.0 mL sample in a microtiter well (24 wells). Make four replicates of 1.0 mL each. Repeat the procedure for all six dilutions. Incubate plate at 18 °C in dark. 7. Observe microtiter plates at 7, 14, and 21 days using phase contrast microscope (40X). 8. Protists are considered to be mycophagous, if ingestion of yeast cells or fungal material is observed or if fungal spores or yeast cells are observed in their cytoplasm [17]. 9. The spores can be fluorescently labeled, and the fluorescencebased survival assay confirmed the fungivore nature of the protists. 3.2 Enumeration of Mycophagous Protists

1. Quantitative work on mycophagous protozoa is very less. Mycophagous amoebae can be enumerated as suggested by Ekelund [3] by the most probable number (MPN) method on agar plates baited with specific fungi [18]. 2. Mycophagous ciliates and testate amoebae actively swimming can be counted directly with counting chamber [19], while number of mycophagous flagellates can be enumerated with method of Hekman et al. [20], using the MPN-method of Darbyshire et al. [21] with Drechmeria coniospora as a food source in microtiter plates. 3. Growth of fungivorous amoeba Protostelium aurantium can be determined by trophozoites inoculated at the center of the plate (day 1) with streaks of different yeasts on low-nutrient agar plates. Plates were examined daily for the growth of P. aurantium and clearing of the yeast over a period of time of 20 days [15].

4

Notes 1. After adding antibiotic to microtiter plate, protection against bacterial colonies can be only for 2 weeks; thereafter either add more antibiotics or transfer to fresh medium. 2. Due care should be taken while performing all isolation steps to avoid contamination especially with antibiotic-resistant strains.

Acknowledgment Authors ADT and BVM are grateful to Principal, Bajaj College of Science, Wardha, for the encouragement and support.

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References 1. Gao Z, Karlsson I, Geisen S, Kowalchuk G, Jousset A (2019) Protists: puppet masters of the rhizosphere microbiome. Trends Plant Sci 24:165–176 2. Petz W, Foissner W, Wirnsberger E, Krautgartner WD, Adam H (1986) Mycophagy, a new feeding strategy in autochthonous soil ciliates. Naturwissenschaften 73:560–562 3. Ekelund F (1998) Enumeration and abundance of mycophagous protozoa in soil, with special emphasis on heterotrophic flagellates. Soil Biol Biochem 30:1343–1347 4. Foissner W, Didier P (1983) Nahrungsaufnahme, Lebenszyklus und Morphogenese von Pseudoplatyophrya nana (KAHL, 1926) (Ciliophora, Colpodida). Protistologica 19: 103–109 5. Flavin M, O’Kelly CJ, Nerad TA, Wilkinson G (2000) Cholamonas cyrtodiopsidis gen. n., sp. n.(Cercomonadida), an endocommensal, mycophagous heterotrophic flagellate with doubled kinetid. Acta Protozool 39:51–60 6. Wilkinson DM, Mitchell EA (2010) Testate amoebae and nutrient cycling with particular reference to soils. Geomicrobiol J 27:520–533 7. Page FC (1988) A new key to freshwater and soil Gymnamoebae. Freshwater Biol. Ass, Ambleside 8. Berthold A, Palzenberger M (1995) Comparison between direct counts of active soil ciliates (Protozoa) and most probable number estimates obtained by Singh’s dilution culture method. Biol Fertil Soils 19:348–356 9. Old KM, Darbyshire JF (1978) Soil fungi as food for giant amoebae. Soil Biol Biochem 10: 93–100 10. Geisen S, Rosengarten J, Koller R, Mulder C, Urich T, Bonkowski M (2015) Pack hunting by a common soil amoeba on nematodes. Environ Microbiol 17:4538–4546 11. Long JJ, Jahn CE, Sa´nchez-Hidalgo A, Wheat W, Jackson M, Gonzalez-Juarrero M, Leach JE (2018) Interactions of free-living amoebae with rice bacterial pathogens Xanthomonas oryzae pathovars oryzae and oryzicola. PLoS ONE 13:e0202941

12. Hibbing ME, Fuqua C, Parsek MR, Peterson SB (2010) Bacterial competition: surviving and thriving in the microbial jungle. Nat Rev Microbiol 8:15–25 13. Frey-Klett P, Burlinson P, Deveau A, Barret M, Tarkka M, Sarniguet A (2011) Bacterial-fungal interactions: hyphens between agricultural, clinical, environmental, and food microbiologists. Microbiol Mol Biol Rev 75:583–609 14. Foissner W (1999) Description of two new, mycophagous soil ciliates (Ciliophora, Colpodea): Fungiphrya strobli ng, n. sp. and Grossglockneria ovata n. sp. J Eukaryot Microbiol 46:34–42 15. Radosa S, Ferling I, Sprague JL, Westermann M, Hillmann F (2019) The different morphologies of yeast and filamentous fungi trigger distinct killing and feeding mechanisms in a fungivorous amoeba. Environ Microbiol 21:1809–1820 16. Geisen S, Koller R, Hu¨nninghaus M, Dumack K, Urich T, Bonkowski M (2016) The soil food web revisited: diverse and widespread mycophagous soil protists. Soil Biol Biochem 94:10–18 17. Scho¨nborn W, Page FC (1988) A new key to freshwater and soil gymnamoebae, with instructions for culture. Freshwater Biological Association, Ambleside 122 S., 55 Abb., zT als Tafeln Brosch 18. Gupta VVSR, Germida JJ (1988) Populations of predatory protozoa in field soils after 5 years of elemental S fertilizer application. Soil Biol Biochem 6:787–791 19. Aescht E, Foissner W (1996) Microfauna. In: Methods in soil biology. Springer, Berlin/Heidelberg, pp 316–337 20. Hekman WE, Van den Boogert PJ, Zwart KB (1992) The physiology and ecology of a novel, obligate mycophagous flagellate. FEMS Microbiol Lett 86:255–265 21. Darbyshire JF, MP G, Rhe I (1974) A rapid micromethod for estimating bacterial and protozoan populations in soil. Revue d’Ecologie et de Biologie du Sol 11:465–475

Part IV Enrichment of Soil Protists on Laboratory Media

Chapter 15 Media Used for Enrichment of Soil Protists Komal A. Chandarana and Natarajan Amaresan Abstract The enrichment of protists highly depends upon the chemical composition of the synthetic medium next to the temperature. It affects growth and reproduction of protists. Therefore, to study the basic life traits such as growth rate and/or interspecies interaction coefficient of different species, standardized media should be used. Many types of media have been used to enrich protists which can be classified as chemically welldefined media and less-defined organic media. Here, we described the five different media which are commonly used to enrich different types of protists. Key words Enrichment media, Life traits, Growth rate, Well-defined media, Organic media

1

Introduction Majority of microcosm studies of protists keep protists in natural freshwater-based medium containing nutrients and sometimes bacteria as food source. The composition of that medium, i.e., nutrient content, pH, and availability of bacterial food, has consequences on enrichment, performance, and evolution of particular protist species. Therefore, during experiments, media composition is adjusted to mimic the environmental conditions from where the species has been isolated. There are a large number of enrichment media widely used for protists, and many types of media have already been described and used [1]. Additionally, manuals for making media commonly are available at different web pages such as http://web. biosci.utexas.edu/utex/media.aspx (UTEX Culture Collection of Algae, University of Texas, Austin), http://www.ccap.ac.uk/ media/pdfrecipes.htm (Culture Collection of Algae and Protozoa, Scottish Marine Institute OBAN, Argyll), and http://tetrahymena. vet.cornell.edu/recipes.php (Tetrahymena Stock Center, University of Cornell, Ithaca) which summarize wider range of media recipes. However, here we only described the five different media which are commonly used and are suitable for a range of different

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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protists to be enriched: proteose peptone medium, protozoan pellet medium, wheatgrass medium (0.1% w/v), yeast-beef agar (YBA) medium and peptone yeast clucose (PYG) agar medium. The former three media are simple less-defined organic media and later two are chemically well-defined media commonly used for laboratory studies of soil protists.

2

Materials

2.1 Basic Requirements

2.2

Media

2.2.1 Proteose Peptone Medium

Graduated beakers and measuring cylinders, micropipettes (range from 0.1 to 10 mL), Erlenmeyer flasks, screw cap autoclavable glass bottles, cotton plug or aluminum foil, spatula, stirring rods 1. Proteose peptone (available through retailers, e.g., Fisher Scientific). 2. 10 μM FeCl3 (prepare it by dissolving 270 mg FeCl3.6H2O in 10 mL of distilled water. 3. 0.2% yeast extract (optional). 4. Bristol medium (use ready-made or make it by dissolving the following ingredients in 1 L distilled water).

2.2.2 Protozoan Pellet Medium

1. Tap water or Chalkley’s solution 2. Protozoan pellet (supplied by standard biological supply company). 3. Protozoan pellets are made of dried and compressed organic material alfalfa.

2.2.3 Wheatgrass Medium (0.1% w/v)

1. Wheatgrass powder (free of pesticidal residues, use organic grass) 2. Distilled water

2.2.4 Yeast-Beef Agar Medium (YBA)

1. Yeast extract (0.5 g) 2. Beef extract (0.5 g) 3. Agar (1.5 g) 4. Distilled water (100 mL)

2.2.5 Peptone Yeast Glucose (PYG) Agar Medium

1. Proteose peptone (10 g) 2. Yeast extract (5.0 g) 3. Agar (20 g) 4. Dextrose (20 g) 5. Distilled water (1000 mL)

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Methods

3.1 Proteose Peptone Medium

Generally, 1 or 2% proteose peptone medium is used which is rich enough to promote high cell density of protists. It is basically the modified Bristol’s medium. This medium can be used for axenic cultivation of ciliate species, especially Tetrahymena sp. [2]. The following procedure should be followed to get 1 L of proteose peptone medium [3]. 1. Fill 900 mL of ready-made Bristol’s medium into the beaker with a minimum volume of 1.5 L. 2. For 1% proteose peptone medium, add 10 mL of proteose peptone to the beaker. 3. Add 100 μL FeCl3 solution. 4. Add 0.2% yeast extract to it (optional). 5. Bring total volume to 1 L by adding Bristol medium. 6. Fill the solution in autoclavable glass bottle and autoclave the medium at 121 °C for 15–20 min (see Note 1).

3.2 Protozoan Pellet Medium

This is the commonly used less-defined medium which is suitable for many protists species. Generally, it is used for bacterized cultures. The content of this medium is not well defined because protozoan pellets are made of dried and compressed organic material. 1. Fill 1 L of distilled tap water or ready-made Chalkley’s medium into autoclavable glass bottle or flask. 2. Add 0.44 g grind protozoan pellet to it. 3. Cover the bottle or flask and autoclave the medium at 121 °C for 15–20 min (see Note 1). 4. Physiochemical description of protozoan pellet medium made in tap water is given in Table 1 (see Note 2).

3.3 Wheatgrass Medium (0.1% w/v)

This is the general medium which can be used to culture and enrich many protistan species. Wheat seeds can also be used and added to the standard solutions in order to provide slow release of nutrients to support the growth of bacteria and protists but are less standardized. 1. Add 1 g of wheatgrass powder to 1 L distilled water into a flask (see Note 3). 2. Bring the solution to boil and let it infused for 2 min. 3. Let it be settled down and cool for an hour. 4. Filter it with several layers of cheesecloth and discard grass residues.

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Table 1 Physiochemical description of protozoan pellet medium Component

Value

Dissolved organic carbon (mg/L)

259.6

Total organic carbon (mg/L)

407

Dissolved nitrogen (mg/L)

24.9

Total nitrogen (mg/L)

33.7

Chloride (mg/L)

72.4

Nitrate (mg/L)

10.8

Sulfate (mg/L)

101.2

Organic phosphorous (μg/L)

225

Dissolved phosphorous (μg/L)

1216

Total phosphorous (μg/L)

2660

Sodium (mg/L)

42.4

Potassium (mg/L)

54.0

Calcium (mg/L)

189

Magnesium (mg/L)

45.8

Ammonium (μg/L)

1501

Nitrite (μg/L)

7.8

Manganese (μg/L)

8.7

Conductivity (μS/cm 20 °C)

1424

Alkalinity (mmol/L)

10.8

Total hardness (nmol/L)

6.9

Silicic acid (mg/L)

137.4

5. Adjust desired pH with 1 N NaOH or 1 N HCl. 6. Dispense medium into autoclavable glass bottle, and autoclave it at 121 °C for 15–20 min (see Note 1). 3.4 Yeast-Beef Agar Medium (YBA)

1. Add 0.5 g of yeast extract powder and 0.5 g of beef extract powder to 100 mL distilled water and stir well. 2. Add 1.5 g agar powder to it. 3. Dispense medium into an Erlenmeyer flask, and autoclave it at 121 °C for 15–20 min (see Notes 4 and 5).

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3.5 Peptone Yeast Glucose (PYG) Agar Medium

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1. Add 10 g of proteose peptone and 5 g of yeast extract powder to 800 mL distilled water, and stir well. 2. Adjust pH to 6.5–7.0 and make the volume up to 900 mL. 3. Add 20 g of agar powder to it. 4. Dispense medium into an Erlenmeyer flask and autoclave it at 121 °C for 15–20 min. 5. Separately, add 20 g of dextrose to another 100 mL of distilled water, dissolve it properly, and filter sterilize it (see Note 6). 6. Add this solution to above autoclaved medium and pour the plates (see Notes 4 and 5).

4

Notes 1. Medium must be cooled down to the normal temperature (around 20 °C) before use. 2. The physiochemical description may vary according to local water and type of protozoan pellet used. 3. Standard soil extract (SSE) can be used instead of distilled water. SSE can be prepared by dissolving 300 g of soil in 1 L of distilled water and stirring well for half an hour. After settling of large soil particles, filter it with cheesecloth and dispense into screw cap bottle and autoclave it. 4. Avoid excessive heating and prolonged sterilization to avoid accumulation of toxic denatured protein derivatives. 5. Plating should be done before cooling down of the medium. 6. Do not autoclave dextrose; otherwise media will turn to brown color because of churning of sugar due to excessive heat of autoclaving.

References 1. Lee JJ, Soldo AT (1992) Protocols in protozoology. Society of Protozoologists, Lawrence 2. Cassidy-Hanley DM (2012) Tetrahymena in the laboratory: strain resources, methods for culture, maintenance, and storage. Methods Cell Biol Acad 109:237–276

3. Asai DJ, Forney JD (2000) Tetrahymena thermophila. Academic, San Diego

Chapter 16 Enrichment of Naked Amoebae Species Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Abstract In spite of various studies on naked amoebae species across the world, the diversity of naked amoebae in various habitats is still poorly studied. The use of appropriate cultivation media and methods is preferred to study the species spectra and ecology in soil and freshwater samples in order to cultivate a broad range of amoebae species. For recovery of the species diversity of naked amoebae, efficiency of cultivation media plays a key role where usually six enrichment media – two liquid and four agar media – are preferred. The enrichment media described here include Prescott-James (PJ), grass-seed infusion (GS), grass-seed agar (GSA), GSA with an overlay of water (GSA + W), non-nutrient agar (NNA), and NNA with an overlay of water (NNA + W). To study the ecology and diversity of naked amoebae from freshwater samples, grassseed infusions were found to be the most effective media, and its preparation method was also found to be easy. Key words Enrichment media, Naked amoebae, Species recovery, Environmental samples

1

Introduction For identification of the naked amoebae diversity to the species level via electron microscopy, light microscopy as well as for molecular analyses in taxonomic studies and high number of specimens are needed [1]. This could be possible by the use of several types of enrichment media used for recovery of the naked amoebae. To support the growth of organisms accompanying amoebae in environmental samples, infusions were the first real culture media used for naked amoebae [2]. A small amount of gelatin or agar can be added to convert these liquid media into a solid form [3]. Some authors proposed the modified culture media with addition of the several nutrients like lettuce extract, beef bouillon, and peptone for obtaining denser cultures [4]. The most commonly used medium for amoebae is found to be the non-nutrient agar medium proposed by Severtzoff [5].

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Amoebae usually feed on living organisms, and bacteria were found to be the most common food for amoebae. To grow such cultures of amoebae feeding on bacteria, media composed of low content of dissolved nutrients are preferred so that the bacteria will not overgrow the amoebae [5]. The media for cultures containing bacteria should not contain rich nutrients; otherwise the bacteria will overgrow compared to the amebae culture. Soil and freshwater amoebae can grow on non-nutrient agar streaked with bacteria such as Escherichia coli grown on ordinary bacteriological nutrient agar. In the case of axenic culture, media containing the dissolved nutrients such as glucose, serum, yeast extract, malt extract, and proteose peptone serve as the only food for amoebae [6]. This book chapter only describes several types of bacterized enrichment media used for recovery of naked amoebae from natural environment.

2

Materials

2.1 Plasticwares and Other Requirements

1. Sample (soil/freshwater) from which amoebae can be recovered. 2. Light microscope/inverted microscope. 3. Glasswares such as pipette, microcapillary, Petri dishes, glass bottles, etc.

2.2

Media

1. Prescott-James (PJ) medium with two rice grains per dish: Prepare it as NNA, but also add Cerophyll/Prescott’s and James’ infusion (CP infusion) for the amoeba saline (AS). AS would be as useful as PJ in making this agar. 2. Grass-seed infusion (GS): Put 2 g of millet grains in 1 L of sterile tap water. Boil the grains in water for 10 min, and then dispense into the Petri dishes at two grains per dish. Prepare this media 1 day before inoculation. 3. Grass-seed agar (GSA): Prepare this media using 1 L of GS and 15 g of non-nutrient agar. Boil it (see Note 1). 4. GSA with an overlay of water from the sample (GSA + W). 5. Non-nutrient agar (NNA): To prepare this media, add 1 L of Neff’s amoeba saline (AS) and 15 g of non-nutrient agar. Boil it. 6. NNA with an overlay of water from the sample (NNA + W).

3

Methods

3.1 Preparation of Media and Inoculation

1. Collect the samples (soil or freshwater) from natural habitats using sterile equipment in sterile glass vessels so as to avoid contamination with cyst-forming amoebae and other organisms from the collecting equipment (see Note 2).

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2. Immediately after transporting to the laboratory, examine the samples microscopically to detect any amoebae with an inverted microscope. 3. Prepare the media used to enrich the naked amoebae (PrescottJames (PJ), grass-seed infusion (GS), grass-seed agar (GSA), GSA with an overlay of water (GSA + W), non-nutrient agar (NNA), and NNA with an overlay of water (NNA + W)). 4. After preparation, autoclave the media using an autoclave at 121 °C for 15 min, and then pour the media into sterile Petri dishes to a depth of 3 or 4 mm. The prepared agar plates can be stored for a week before use if kept in clean conditions to minimize contamination by mites or fungi (see Note 3). 5. Inoculate 0.2 mL of sediment/freshwater sample into the appropriate medium, according to the amoeba sought. 6. Incubate the cultures at 22–37 °C in indirect light for 6–7 days (see Note 4). 7. To enrich the amoebae, subculture the sample into the appropriate medium according to the amoeba which is sought. For subculturing, viable cell count should be numerous [6]. 3.2 Enrichment of Naked Amoebae on Liquid Media

1. If amoebae are grown on liquid media, transfer the amoebae in the subsequent media with the help of a pipette or capillary. If amoebae are grown in protein-rich medium, then firstly agitate the culture with a pipette, and then transfer a pipette-full to the new medium. For non-axenic cultures in liquid, pour fresh medium into a sterile Petri dish in equal quantity to the parent culture. 2. Transfer the entire parent culture into a new dish with a pipette, mix it well with the fresh medium, and then transfer the half of mixed liquid back into the old Petri dish. This may rejuvenate the parent culture and provide a large inoculum for the daughter culture [7].

3.3 Enrichment of Naked Amoebae on Solid Agar Media

1. For enrichment of the amoebae grown on agar cultures, transfer an agar block from the parent culture to a fresh agar surface. If the medium used is NNA, streak the bacteria such as E. coli as a single streak on the fresh surface before inoculating the amoebae, and then place the agar block at one end. This prolongs the culture life as the amoebae will reach only to the other end of the streak before exhausting their food supply. 2. If few amoebae are present or amoebae multiply slowly on agar, then wash the agar surface with 1 mL of liquid media. Pipette out this suspension onto a fresh surface, and also tilt the plate to distribute the liquid in a proper manner. This is done to

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prevent the bacteria from overgrowing the amoebae and to allow development of several foci of growth especially when the media is enriched with the nutrients (GSA or PJ). 3. After inoculation in the agar plates, incubate it either in normal or in inverted position (see Note 5). 3.4 Enrichment of Naked Amoebae Formed as Cysts

1. For enrichment of the amoebae that form cysts, wash the agar surface of the mixed culture with 0.5–1 mL of AS or PJ. With the help of a bacteriological loop, streak this suspension onto NNA, and then find out the isolated cyst as cysts can be seen easily and less likely to be damaged than active amoebae. 2. Cut a 1–2 mm block of agar bearing a cyst with a sterile fine scalpel, and put it on a fresh surface of the appropriate agar. This block of agar contains some bacteria also along with the cyst from natural habitats, these bacteria will multiply when agar block is inoculated in medium such as GSA which provides food for the amoebae. 3. Incubate the amoebae cultures in liquid or solid media for 6–7 days for enrichment of the amoebal count. 4. Visualize them under an inverted microscope so as to note whether the amoebal culture was enriched or not in terms of count and preserve them via cryopreservation [7].

4

Notes 1. While pouring the grass-seed agar into the Petri dish, try to distribute the same number of seeds evenly in each Petri dish. All the grass seeds from which the infusion was made should be left in the agar. 2. Sample collection should be handled with sterile equipment until the sample is inoculated into the appropriate media. 3. Shelves and cupboards should be kept clean to store the media plates to minimize the possibility of contamination with mites. 4. Do not incubate the cultures in direct sunlight especially the liquid cultures other than axenic cultures should be kept in the dark to prevent growth or algal contaminants. 5. The best way to incubate the plates is the normal position, at least for the first few days, so as to avoid the drop off of the agar block bearing the inoculum or to ensure that the suspension does not run down onto the lid of the plate.

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References 1. Mrva M, Garajova´ M (2018) The efficiency of cultivation media in recovering naked lobose amoebae from freshwater environments. Zool Stud 57 2. Monica Taylor SND (1925) Amoeba proteus: some new observations on its nucleus, lifehistory, and culture. Q J Microsc Sci 69:119 3. Jollos V (1917) Untersuchungen zur orphologie der Amo¨benteilung. Arch Protistenkd 37:229– 275

4. Doflein F (1953) Lehrbuch der protozoenkunde. 6. aufl. von E. reichenow. VEB G Fischer, Jena 5. Severtzoff LB (1924) Method of counting, culture medium and pure cultures of soil amoebae. Centralb F Bakteriol Orig 92:151–158 6. Smirnov AV, Chao E, Nassonova ES (2011) Cavalier-Smith T. 2011b. A revised classification of naked lobose amoebae. Protist 162:545–570 7. Kalinina LV, Page FC (1992) Culture and preservation of naked amoebae. Acta Protozool 31: 115–126

Chapter 17 Enrichment of Ciliates and Flagellates Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Abstract Ciliates and flagellates are the group of protozoans, found abundant in agricultural and forest soil, as well as in freshwater, coastal, and marine water sediments. They play an important role as bacterial consumers and as fungi consumers to some extent leading to organic soil nitrogen mineralization which was then taken by the plants. This bacterial consumption by ciliates and flagellates enhances the nutrient cycles and the energy flow to the plants, animals, and microorganisms as well and, therefore, is considered as effective bioindicators of soil quality. Ciliates and flagellates also play an essential role in sustainable agricultural development as they can interact with native bacteria and produce negative effect on plant diseases by stimulating antibiotic and secondary metabolite production through enhancing the growth of plant growth-promoting bacterial species. Therefore, to recover the species diversity of ciliates and flagellates, cultivation media plays a key role, where hay infusion, proteose peptone (PP), skim milk-based media, chemically defined media (CDM), and alfalfa enrichment medium are generally preferred. Key words Enrichment media, Ciliates, Flagellates, Bioindicators, Hay infusion

1

Introduction Ciliated protozoa are dikaryotic unicellular organisms, and flagellates are organisms with one or more whip-like appendages called flagella. These groups of protozoans can be easily identified based on their special characteristics of nuclei, flagella, infraciliature, and locomotion. Their role as bioindicators has been demonstrated indicating that the ciliates and flagellates had a strong correlation with soil physical and chemical parameters [1]. Apart from that, they are also useful for plants, as they produce negative effects on plant diseases by interacting with native bacteria, i.e., the ciliates and flagellates may stimulate the growth of plant growthpromoting bacterial species which, in turn, may stimulate the antibiotic and secondary metabolite production and, hence, improve the bacterial ability to suppress plant diseases [2]. This led to the sustainable agricultural development [2].

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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There are around 3000 free-living ciliated species, and it has been estimated that this number may reach up to 30,000 species [3]. Free-living ciliates and flagellates are ubiquitous in nature and can be successfully cultured in the laboratory. Moreover, there are many new ciliate and flagellate species which are either described or redescribed with species being undiscovered [4]. Hence, there is an urge need to conduct a more intensive research on the diversity of ciliates and flagellates. The most commonly used media for laboratory culture of ciliates and flagellates such as Tetrahymena and Paramecium includes proteose peptone (PP)-based media, hay infusion, skim milk-based media, alfalfa medium, and chemically defined media (CDM) [5]. This book chapter, therefore, provides basic information for the several types of enrichment media used for the isolation and recovery of ciliates and flagellates.

2

Materials

2.1 Basic Requirements

1. Sample (soil/freshwater) from which ciliates and flagellates can be isolated or recovered. 2. Light microscope/inverted microscope/stereoscope 3. Glasswares such as microcapillary, pipette, Petri dishes, glass bottles, etc.

2.2

Media

1. Hay infusion: Ciliates and flagellates are originally cultured in bacterized hay or vegetable matter infusions. To prepare the hay infusion, boil 10 g chopped Timothy hay (other types of hay can also be used) in 1 L of spring water for half an hour. After boiling, filter this mixture through several layers of cheesecloth, and allow the solution to cool down. Then add a pinch of black soil and two drops of 1 M NaOH to the cooled mixture. Now, pour this liquid medium to each culture dish (approximately, 200 mL), and allow to stand uncovered for 24 h. After that, add either two cooked and crushed wheat seeds or 1–2 g of dried and crushed lettuce to each dish (see Note 1). This media is generally used for the recovery of ciliates and flagellates [6]. 2. Alfalfa medium: To prepare this media, firstly, prepare 10× Chalkey’s stock solution where 0.06 g CaCl2, 1.00 g NaCl, and 0.04 g KCl were dissolved in 1 L distilled water. Dilute by a factor of 10 to make it as 1× solution (see Note 2). Now, add one alfalfa in one liter of boiling 1× Chalkey’s solution (see Note 3), and allow the alfalfa to soak until it forms a sediment on the bottom of the container. This sediment can be used as a source of food for the bacteria present along with the ciliates and flagellates. Add ten boiled wheat seeds to 750 mL of cooled alfalfa medium for culturing ciliates and flagellates.

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Inoculate 100 mL of ciliate and flagellate culture in this medium for enrichment of ciliates and flagellates. Incubate the culture for around 3 weeks for reaching to its peak [6]. 3. Proteose peptone (PP)-based media: PP media is a rich nutrient media comprising of 2% proteose peptone and 10 μM FeCl3 or 90 μM sequestrene Fe-EDTA. Generally, growth in all PP media is limited due to the limited Fe, but the Fe need was met by supplementation with Fe salts (FeCl3) at a low concentration of 10 μM or chelated Fe salts like Fe-EDTA or sequestrene. Yeast extract also contains Fe, but yeast extract with an additional Fe supports the growth of ciliates and flagellates. PP media with liver extract does not require Fe. If ferric or ferrous chloride is added prior to autoclaving in the media, it may produce an iron precipitate (see Note 4). To avoid or to prevent precipitation, filter-sterilize a concentrated Fe solution separately, and then add it to the autoclaved media immediately when it gets cooled or prior to its use. Filter sterilization of any PP media may affect the growth of the culture as some particulate matters are necessary for the formation of food vacuoles. Therefore, as a substitute, Fe-EDTA is used as it does not form a precipitate when added to the media before autoclaving. The prepared PP media is then autoclaved at 121 °C and 15 psi for about 30 min (see Note 5) [7]. 4. Skimmed milk media: This media comprises of 2% skimmed milk, 0.5% yeast extract, 0.1% ferrous sulfate chelate solution, and 1% glucose solution. For large-scale cultivation of ciliates like Tetrahymena, less expensive media, i.e., skim milk-based media can be used. During conditions of high cell density fermentation and cell retention in a bioreactor, the culture of Tetrahymena at cell density of more than 2.2 × 107 cells/mL, equivalent to 48 g dry weight, was supported by skim milkbased medium. It has also been revealed that for supporting growth up to ~3 × 106 cells/mL in batch fermenters, diluted skim milk-based medium, MYE medium comprising of 1% skim milk, and 1% yeast extract have been used [7]. 5. Chemically defined media (CDM): The amino acids, vitamins, purines and pyrimidines, and the solutions of chemicals were used to prepare the CDM. Prepare the amino acids as a dry mixture, and prepare the vitamins with mannitol as triturate and also the chemicals and media that had been stored with volatile preservative with twice glass-distilled water. The composition of the CDM thereby adjusting the constituents’ concentration has been mentioned in Table 1. pH of the media was adjusted to 6.5 before autoclaving, and then the medium was autoclaved at 15 lb/sq in for 30 min [7].

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Table 1 Composition of chemically defined media (CDM) Amino acids

Conc. (mg/100 mL)

Vitamins

Conc. (μg/100 mL)

L-arginine HCl

45.0

Thiamin HCl

500.0

L-tyrosine

2.0

Inositol

1000.0

L-histidine, base

32.5

Nicotinamide

100.0

DL-serine

35.0

p-Aminobenzoic acid

10.0

DL-isoleucine

25.0

Ca-d-pantothenate

100.0

L-proline

2.0

Choline chloride

500.0

L-leucine

25.0

Pyridoxamine 2 HCl

100.0

Glycine

25.0

Pyridoxine

100.0

L-lysine 2 HCl

20.0

Riboflavin

50.0

L-glutamic acid

50.0

Biotin

0.5

DL-methionine

9.0

Folic acid

5.0

L-cystine

2.5

B12

0.1

DL-phenylalanine

32.0

Mannitol, as triturate

2400.0

DL-aspartic acid

25.0

Salts

(mg/100 mL)

DL-threonine

20.0

MgSO4.7H2O

20.0

DL-alanine

35.0

Na2HPO4

20.0

DL-tryptophan

25.0

KH2PO4

20.0

DL-valine

17.5

CaCl2

Purines and pyrimidines

(mg/100 mL)

Uracil

0.1

Guanylic acid

1.0

Adenylic acid

0.5

Cytidylic acid

0.5

Trace metal solution

1.0 *

1 mL/100 mL

*

1 mL/100 mL trace metal solution provides metal ion concentration in μg: Fe 100, Mn 65, Zn 22.6, Mo 20, Cu 0.13, Co 0.63, EDTA Na2 1000. pH of medium was adjusted to 6.5

3

Methods 1. Collect the soil or freshwater samples from which ciliates and flagellates are to be recovered using sterile equipment in order to avoid contamination from the collecting equipment. 2. Immediately transfer the samples to the laboratory, and examine the samples microscopically to detect ciliates and flagellates with an inverted microscope or stereoscope.

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3. Prepare the media to enrich the ciliates and flagellates such as proteose peptone (PP) based media, hay infusion, skim milkbased media, alfalfa medium, and chemically defined media (CDM). 4. Autoclave the media using an autoclave as mentioned above, and then inoculate 0.2 mL of sediment/freshwater sample into the appropriate medium, to recover the ciliates and flagellates. 5. Incubate the cultures at 22–37 °C for 3 weeks to recover the more no. of ciliates and flagellates (see Note 6). 6. To enrich the culture of ciliates and flagellates, subculture the sample into 200 mL of the appropriate fresh medium, and add 5–6 mL of the ciliate or flagellate culture as inoculum. For subculturing, viable cell count should be numerous. 7. Monitor the cultures of ciliates and flagellates regularly using a stereoscope [6].

4

Notes 1. Before adding wheat seeds or crushed lettuce to the media, boil wheat seeds or crushed lettuce in a few mL of water for 15 min, and also keep in mind that the lettuce leaves should be dried slowly in an oven, grounded with mortar and pestle and the dried lettuce stored in a tightly sealed container for future use. 2. To make 1× Chalkey’s solution from 10× stock solution, dilute 100 mL to 1 L with distilled water. 3. Purchase alfalfa pills from garden shops, farm supply stores, and health food stores. 4. Fe precipitate does not affect the growth but may interfere with some downstream operations such as electronic cell counting or cell collection by high-speed centrifugation. 5. The ability of the media to support optimal growth of ciliates and flagellates may get decrease due to excessive autoclaving. 6. Incubate the plates in the normal position to avoid the drop off of the agar block bearing the inoculum or to ensure that the suspension does not run down onto the lid of the plate.

References 1. Madoni P (2011) Protozoa in wastewater treatment processes: a minireview. Ital J Zool 78:3– 11 2. Geisen S, Mitchell EA, Adl S, Bonkowski M, Dunthorn M, Ekelund F, Lara E (2018) Soil

protists: a fertile frontier in soil biology research. FEMS Microbiol Rev 42:293–323 3. Finlay BJ, Esteban GF (1998) Freshwater protozoa: biodiversity and ecological function. Biodivers Conserv 7:1163–1186

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4. Foissner W (2015) Terrestrial and semiterrestrial ciliates (Protozoa, Ciliophora) from Venezuela and Gala´pagos. Denisia Press, Linz, p 35 5. Cassidy-Hanley DM (2012) Tetrahymena in the laboratory: strain resources, methods for culture, maintenance, and storage. Methods Cell Biol Acad 109:237–276

6. Hyman LH (1931) Methods of securing and cultivating protozoa. II. Paramecium and other ciliates. Trans Am Microsc Soc 50:50–57 7. Adam KM (1959) The growth of Acanthamoeba sp. in a chemically defined medium 519. Microbiology 21:519–529

Chapter 18 Enrichment of Mycophagous Protists Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Abstract Soil protists are considered to be bacterivorous and fungivorous where bacterial energy channel in food web is controlled by bacterivorous nematodes and the fungal energy channel is controlled by arthropods and mycophagous nematodes. Most of the soil biologists accept this perspective; however, some taxonomists challenged the functional studies and revealed a range of mycophagous protists. Mycophagy among soil protists is common and is of ecological importance in terrestrial ecosystems. To increase the knowledge on the functional importance of mycophagous protists, there is an urge need of cultivating mycophagous protists. Therefore, for isolating and for enrichment of mycophagous protists, Neff’s modified amoeba saline (NMAS) and malt extract agar (MEA) are used. Fungal suspension as a feeding material for mycophagous protists is also needed. Key words Enrichment media, Mycophagous protists, Fungivorous, Neff’s modified amoeba saline, Malt extract agar

1

Introduction Protists are of marginal importance during fungal energy channel especially since mycophagous nematodes and microarthropods are found to be predominant consumers [1]. Protist taxonomists found that facultative and obligate mycophagous protists diversity are more common in soils such as ciliates belonging to the family Grossglockneriidae (obligatory mycophagous), soil eukaryotes such as Thecamoeba spp., vampyrellid amoebae, few testae amoebae, and eumycetozoans (facultative mycophagous protists – omnivores) [2]. It is a well-known fact that mycophagous protists have similar biomasses of those of bacterivorous protists, but still the detailed study about the functional diversity of mycophagous protists is limited in terrestrial ecosystem [3]. Several experiments revealed that a variety of cultivable mycophagous protists and closely related protists feed bacterial prey, but mycophagous pro-

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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tists also feed on fungi thereby leading to shifts of fungal community of soils [4]. Some former studies hypothesized that yeasts are preferred as a food material for protists, whereas hyphae-forming fungi is less preferred as a suitable prey [5]. Mycophagy among soil protists is common, and to evaluate the ecological and functional diversity of mycophagous soil protists, mycophagous protists are cultivated, and this can be made possible by the use of enrichment media. Therefore, this book chapter deals with the enrichment media preferred for recovery of mycophagous protists where NMAS and MEA are preferred with fungal suspension as a feeding material.

2

Materials

2.1 Basic Requirements

1. Environmental samples such as soil for recovery of mycophagous protists 2. 24 microwell-plate, orbital shaker 3. Centrifuge 4. Light microscope/inverted microscope

microscope/phase

contrast

5. Glasswares such as pipette, microcapillary, Petri dishes, Parafilm, glass bottles, etc. 6. Fungal inoculum as a feeding material for enrichment of mycophagous protists 2.2

Media

1. Neff’s modified amoeba saline (NMAS): Prepare solution A and solution B. For preparing solution A, 1.20 g of NaCl, 0.04 g MgSO4.7H2O, 1.42 g Na2HPO4, and 1.36 g KH2PO4 were dissolved per 100 mL of distilled water in a glass beaker. Prepare solution B separately by dissolving 0.04 g of CaCl2.2H2O per 100 mL of distilled water. Finally, take another glass beaker and add amoeba saline to it. Mix 10 mL of solution A and 10 mL of solution B to it. Pour 980 mL of distilled water, make the final volume up to 1000 mL, and adjust the solution of this NMAS to 6.9 by using 1N KOH [6]. 2. 1.5% malt extract agar (MEA): To prepare MEA plates, add 1.5% of malt extract powder and 0.5% of non-nutrient agar to it. Autoclave the media at 122 °C for 20 min. Pour the media into the autoclaved Petri plates, and allow it to get solidified [1].

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3

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Methods 1. Collect the soil samples from the environment in a sterile polybags using the sterile equipment (see Note 1). 2. Examine the samples microscopically using a light microscope or an inverted microscope to detect mycophagous protists. 3. For isolation of mycophagous protists, enrichment cultures were established. 4. Weigh 1 g of soil sample, and suspend it in 250 mL of Neff’s modified amoeba saline (NMAS). 5. Shake the suspension on an orbital shaker at 100 rpm for 10 min, and make fourfold dilution with NMAS. 6. Add 20 μL of the suspension to the wells of a 24 multiwell plate. 7. Add a fungal inoculum to each well as a feeding material for enriching the mycophagous protists (e.g., a mixed fungal inoculum of dried Saccharomyces cerevisiae and a Fusarium culmorum could be added to each well, i.e., 80 μL of a 0.4 g/L NMAS solution of dried S. cerevisiae and 160 μL of a F. culmorum with four spores/μL concentration. 8. Seal the 24-well plates using Parafilm, and store them at 15 °C in the dark condition (see Note 2). 9. Incubate the plates for 21 days. 10. Observe the enrichment cultures microscopically after 7 and 21 days of incubation at 100× and 400× magnification of an inverted microscope to examine mycophagous protists, i.e., growth on fungi and ingestion of fungal material. 11. To establish active fungal cultures, MEA media can also be used as enrichment culture of mycophagous protists where MEA plates can be inoculated with 100 μL of F. culmorum spores and hyphae suspension. 12. Record the microphotographs of protists ingesting fungal material [2].

4

Notes 1. Soil samples should be collected with sterile equipment until and unless it is inoculated into the appropriate media in order to maintain sterility. 2. Store the cultures in the dark so as to prevent growth or algal contaminants.

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References 1. Geisen S, Koller R, Huenninghaus M, Dumack K, Urich T, Bonkowski M (2016) The soil food web revisited: diverse and widespread mycophagous soil protists. Soil Biol Biochem 94:10–18 2. Stephenson SL, Feest A (2012) Ecology of soil eumycetozoans. Acta Protozool 51(3) 3. Ekelund F (1998) Enumeration and abundance of mycophagous protozoa in soil, with special emphasis on heterotrophic flagellates. Soil Biol Biochem 30:1343–1347

4. Glu¨cksman E, Bell T, Griffiths RI, Bass D (2010) Closely related protist strains have different grazing impacts on natural bacterial communities. Environ Microbiol 12:3105–3113 5. Allen PG, Dawidowicz EA (1990) Phagocytosis in Acanthamoeba: I. A mannose receptor is responsible for the binding and phagocytosis of yeast. J Cell Physiol 145:508–513 6. Rowbotham TJ (1983) Isolation of Legionella pneumophila from clinical specimens via amoebae, and the interaction of those and other isolates with amoebae. J Clin Pathol 36:978–986

Chapter 19 Mono-axenic Cultivation of Protists Komal A. Chandarana and Natarajan Amaresan Abstract Several research studies require the application of specific bacterial species with protist; however, it is very difficult to eliminate native bacterial population from protistan culture. Application of antibiotics may always not work well. Therefore, we here described the easiest way to prepare mono-axenic culture of protists with specific bacterial species for research studies employing simultaneous applications of bacteria and protists. Key words Research studies, Native bacterial population, Antibiotics, Mono-axenic

1

Introduction Isolation and separation of protists from environmental samples for new research studies always accompanied native bacterial population which may later hinder the results of the studies. Therefore, to remove these bacteria, researchers commonly applied antibiotics such as penicillin/streptomycin, by determining appropriate concentration empirically. Further, axenic cultivation of protist by using high-nutrient media along with antibiotics serves as an alternative method to enrich protists for the studies. However, these antibiotics cannot eliminate all bacterial populations from the protistan culture samples. Moreover, many bacterivorous protists require the presence of bacteria as a food source for its growth and survival, especially amoebae which cannot easily be cultivated axenically [1]. In addition to this, some studies require the simultaneous application of specific bacterial species and protistan species [2–4]. And in such type of experiments, native bacterial population definitely affects the outcome of the study. Therefore, we here described the mono-axenic culture preparation of the protists with specific bacterial species according to the method described

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_19, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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by Chandarana and Amaresan [4]. The protocol is developed in our laboratory by gathering ideas from several previous studies and by experience of work with several soil protists.

2

Materials

2.1 Glasswares and Equipment

1. Petri dishes, wire loop, microcentrifuge tubes, micropipettes, Erlenmeyer flask 2. Microscope, centrifuge

2.2

Media

1. 0.2X nutrient yeast extract agar (NYEA): Weigh 1.6 g nutrient broth (HIMEDIA), 0.4 g yeast extract, 1.0 g glucose, and 20.0 g agar powder. Mix the ingredients in 1.0 L distilled water. Sterilize it by autoclaving at 15 Ibs pressure, 121 °C for 15 min. Pour the media in transparent glass Petri dishes. Do not make thick layer of agar; otherwise it will hinder the observation of microbes under the microscope. 2. Nutrient agar media: Suspend 12.8 g nutrient agar (HIMEDIA) in 100 mL distilled water, and sterilize it by autoclaving. Pour the media in Petri plates. 3. 1% nutrient broth Page’s saline (NB-PAS) agar media: First, make Page’s saline by suspending 0.0403 g of Page’s saline (HIMEDIA) in 100 mL distilled water. Now, add 0.13 g nutrient broth (HIMEDIA) to this solution. Mix well the content. Sterilize it by autoclaving and pour it in Petri dishes.

2.3

Cultures

1. Bacterial cultures (see Note 1) 2. Protist culture

3

Methods

3.1 Bacterial Culture Preparation

1. Take a stock culture of desired bacterial species, and make a fresh culture of it by streaking it on sterile nutrient agar media (see Notes 2 and 3).

3.2 Protist Culture Preparation

1. Take enriched protist growth in microcentrifuge tube, add 1.0 mL Page’s saline to it, and wash the protist cells by repeated pipetting. 2. Centrifuge the content at 1000 rpm for 8 min. 3. Repeat the procedure two to three times by discarding supernatant to reduce the number of protist-accompanied bacterial cell and spent medium.

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Fig. 1 Summary of the experimental design conducted to prepare the mono-axenic culture of protist with specific bacterial species. (Reproduced from ref. 4 with permission from Journal of Applied Microbiology, Oxford University Press) 3.3 Mono-axenic Culture Preparation

1. Streak a fresh culture of selected bacteria as two parallel lines across 0.2X nutrient yeast extract agar (NYEA) medium (Fig. 1) (see Note 4). 2. After 3 h of incubation, inoculate 5 μL (200 cells mL-1) of a protist culture (either trophozoites or cyst or a mixture of active protist and cysts) onto one end of the streaked bacterial line. 3. The second bacterial line will be served as a control to compare the effects of predation activity on bacterial growth. 4. Incubate the Petri dishes for 5 days at room temperature under dark conditions. 5. Observe the plate visually as well as directly under the 10× objective of dissecting microscope. The elimination of confluent bacterial growth along the streaked line or changes in growth characteristics of the bacterial line revealed the migration and predation of protists on that bacterial strain. The presence of protists along the bacterial line ascertained under the microscope will confirm the migration of protists along the selected bacterial growth (see Note 5). 6. After confirming, collect the protist isolate from the second edge of streaked bacterial line (which contained protist with specific bacterial species only) (Fig. 1).

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7. Transfer this collected growth on fresh 1% NB-PAS agar plates. No other bacteria were added as food sources (see Note 6). 8. Repeat the steps 1–7 three to four times to avoid the contamination by protist-accompanied bacteria (other bacteria). However, at step 2, inoculate protist collected from step 7. This repetition of process will gradually eliminate the protistaccompanied bacterial growth. 9. To confirm each specific bacterial species with inoculated protist, streak the collected combined culture onto separate nutrient agar plates. Incubate the plates at appropriate conditions, and observe the bacterial colony characteristics.

4

Notes 1. The commonly recommended strains of bacteria are Escherichia coli, Enterobacter aerogenes, and Klebsiella pneumoniae subsp. Pneumoniae on non-nutrient agar media to prepare mono-protist culture [1]. However, any pure culture of bacteria can be used to prepare mono-axenic culture of protists with that specific bacterial species. The only concern is that the protist must be able to feed on that bacterial species. 2. One can use bacteria-specific media instead of nutrient agar media. Nutrient broth can also be used to make fresh culture of bacterial species. 3. It is recommended to use fresh culture (16–24 h old) of bacteria. 4. Make appropriate distance between two parallel bacterial lines. The bacterial lines should not be too close that protist migrates to second line also. 5. Sometimes the streaked bacterial line becomes mucoidal growth due to bacterial defense against predation pressure, so the observation of protistan growth can be challenging. Especially for the inexperienced, it can be quite difficult to discern protist amidst the dense growth of bacterial flora. 6. Protist in xenic culture may exhibit reduced or inhibited growth if bacterial density becomes excessive. Therefore, to obtain better protistan growth, bacterial densities must be reduced to some extent which can be controlled either by diluting media or by adding antibiotics to media. 1% NB-PAS media serve the low nutrient content which will favor the protistan growth as well as bacterial growth; however, it can help to modulate bacterial growth.

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Acknowledgment This work was supported by the Department of Science and Technology, Science and Engineering Research Board (DST-SERB), New Delhi, India, under an ECRA grant for researchers to NA (ECR/2017/001977). References 1. Nerad TA (ed) (1991) American type culture collection catalogue of protists, 17th edn. American Type Culture Collection, Rockville, p 88 2. Chandarana KA, Pramanik RS, Amaresan N (2022) Interaction between ciliate and plant growth promoting bacteria influences the root structure of rice plants, soil PLFAs and respiration properties. Rhizosphere 21:100466

3. Chandarana KA, Pramanik RS, Amaresan N (2022) Predatory activity of Acanthamoeba sp genotype T4 on different plant growthpromoting bacteria and their combined effect on rice seedling growth. Eur J Protistol 82: 125858 4. Chandarana KA, Amaresan N (2023) Predation pressure regulates plant growth promoting (PGP) attributes of bacterial species. J Appl Microbiol 134:lxad083

Chapter 20 Axenic Cultivation of Soil Protists Sapna Chandwani, Hetvi Naik, and Natarajan Amaresan Abstract Most of the physiological work on the protozoa has been restricted because the organism which need to be investigated cannot be raised in pure culture, i.e., without the presence of any other organisms. Due to difficulties in obtaining axenic cultures (pure culture), biochemical, genetic, physiological, and taxonomic studies have been limited. Axenic cultivation of soil protists is also required for accurate identification of the protozoan species. Protein-rich liquid media are generally used for cultivation of soil protists axenically. Key words Axenic culture, Soil protists, Protein-rich liquid media, Pure culture

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Introduction The production and maintenance of protist cultures without the presence of other organisms where the species is being cultured are termed axenicization. Generally, Acanthamoeba and Naegleria spp. (free-living amoebae) can be grown axenically. Soil protists like amoebae are generally cultivated axenically by using antibiotics like ampicillin, cefoxitin, meropenem, penicillin, and streptomycin in the selective media and liquid solutions used to cultivate the soil protists such as phosphate-buffered soil saline (PBS), 1.5% agar media, standard soil extract (SSE), 0.1% wheat grass medium, yeast-beef agar medium (YBA), Prescott-James medium (PJ), non-nutrient agar (NNA), 1% proteose peptone medium, and grass-seed infusion (GS). These antibiotics may inhibit the bacteria growing along with the protists; however, inhibiting the bacterial growth may not always lead to the complete isolation of soil protists. Compared to other protists, some species of these genera are very difficult to axenicize and some Acanthamoeba strains can never be axenicized [1]. For many biochemical and biophysical investigations, such as in the case of free-living protozoa Tetrahymena and Paramecium, successful axenic cultivation contributed greatly to their usefulness [2]. The axenic cultivation of soil amoebae like

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_20, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Acanthamoeba has made possible the metabolic and electron microscopic studies; however, morphological heterogeneity and long generation times limited the type of experiment that can be carried out [3]. In this book chapter, we presented the information on the axenic cultivation of soil protists so as to use this organism as a model system to study the biochemical, genetic, physiological, molecular, and taxonomic studies.

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Materials

2.1 Glass- and Plasticwares and Other Requirements

1. Soil sample from which protists can be isolated 2. Light microscope/ inverted microscope 3. Glasswares such as pipette, microcapillary, Petri dishes, glass bottles, etc. 4. Antibiotics such as ampicillin, cefoxitin, meropenem, penicillin, and streptomycin

2.2

Media

1. Non-nutrient agar (NNA): To prepare this media, add 1 L of Neff’s amoeba saline (AS) and 15 g of non-nutrient agar. Boil it. 2. Proteose peptone glucose (PPG): To prepare this media, dissolve 10 g proteose peptone and 18 g glucose in 1 L of Neff’s amoeba saline (AS), and then autoclave the media in culture tubes. 3. Chang’s serum/casein/glucose/yeast extract medium (SCGYEM): Dissolve 10 g of isoelectric casein, 2.5 g glucose, 5 g yeast extract, 1.325 g Na2HPO4, 0.8 g KH2PO4, and 100 mL fetal calf serum in 900 mL glass-distilled water. Add 200 μg/mL penicillin and streptomycin if the inoculum contains bacteria. Autoclave the media in culture tubes. 4. Jones’s medium: To prepare this buffered saline, dissolve 2.65 g Na2HPO4.12H2O, 0.41 g KH2PO4, and 7.36 g NaCl in 1 L of glass-distilled water. Adjust the pH to 7.2. Make sure that the medium should contain 850 mL buffered saline, 50 mL horse serum, and 100 mL of 1% yeast extract solution. Sterilize the media and pour it into the sterile tubes. To each tube, add a pinch of sterile rice starch (see Note 1).

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Methods

3.1 Sample Collection and Isolation

1. Collect the soil samples from natural habitats with sterile equipment in sterile glassware in order to avoid contamination from the collecting equipment (see Note 2).

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2. After transporting soil samples to the laboratory, examine the samples containing protozoan species microscopically with an inverted microscope. 3. Inoculate the samples to the selective medium such as phosphate-buffered soil saline (PBS), 1.5% agar media, standard soil extract (SSE), 0.1% wheat grass medium, yeast-beef agar medium (YBA), Prescott-James medium (PJ), non-nutrient agar (NNA), 1% proteose peptone medium, and grass-seed infusion (GS) according to the protozoan species which are sought. For example, proteose peptone medium to grow Tetrahymena spp.; wheat grass medium for culturing protozoa such as rhizopods, choanoflagellates, ciliates and flagellates; Prescott-James medium (PJ); non-nutrient agar (NNA); grass-seed infusion for naked amoebae; 1.5% agar medium; and soil extract medium for Acanthamoeba spp. (refer to Chap. 16: “Enrichment of Naked Amoebae Species” in the present book for media listed in this step). 4. Incubate the cultures at 22–37 °C for 6–7 days for isolating the protozoan species (see Note 3). 5. Enrich the protozoan species by subculturing the samples into the appropriate medium, and for subculturing, viable cell count should be numerous. 6. Along with the protozoa, bacteria or small protists which may serve as food for the protozoa like large amoebae need to be eliminated for cultivating the protists axenically if one has to study the biochemical, genetic, physiological, molecular, and taxonomic studies. 7. To study the protozoan species, axenicization is required [2]. 3.2 Axenic Cultivation of Soil Protists 3.2.1

Using Antibiotics

3.2.2 Migration Method of Axenicization

1. Inoculate the antibiotics such as ampicillin, cefoxitin, meropenem, penicillin, and/or streptomycin into the media used to subculture the protozoan species so as to allow the growth of the protists axenically. 2. Use of antibiotics may inhibit the bacteria to a greater extent but will not always lead to complete axenic cultivation of protists. Therefore, migration method of axenicization is preferred to maintain the purity of the culture [1]. 1. Cultivation of protists axenically is also made possible by inoculating protists such as amoebae onto the solid media containing agar with killed bacteria (sometimes heat-killed), but this migration method of axenicization uses live bacteria and is successful for cultivation of soil protists like Acanthamoeba. 2. Depending on the migration ability of protozoa to a bacteriafree area of an agar media, from where they are taken before and later brought bacteria to that area, axenicization is achieved.

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3. First of all, prepare NNA media, sterilize it using an autoclave, and pour the media into the Petri dish. Allow it to solidify. 4. After solidification of the media, make a crescent-shaped streak of bacteria such as E. coli on one side of the NNA media plate. 5. Inoculate a drop of protist suspension between the bacterial streak and the side of the media plate. 6. The protist will migrate toward the bacteria to feed the bacteria, get multiplied, and then further keep on migrating. 7. The first protist that will migrate 1 cm or more beyond the bacterial streak will be free of bacteria. Immediately, transfer this protist to an agar block and then to liquid medium as soon as possible for the maintenance of axenic culture of protists. 8. Generally, protein-rich liquid media and sometimes Jones’s medium are preferred for axenic cultivation of soil protists. 9. Take a single drop of the protist culture from the top of the liquid medium, and examine the culture under microscope (see Note 4). 10. Incubate the protist culture for a few days, and after incubation, discard the cultures that are highly contaminated with bacterial growth. 11. If protist cells are found and no bacteria are seen under the microscope view, then allow the culture to grow more for a week or two, and then test the culture for purity. 12. Purity of the culture can be tested by subculturing the axenic protist culture to a fresh medium at regular intervals and maintaining them for further study (biochemical, genetic, physiological, molecular, and taxonomic studies) as well as for identification of the protist culture [1, 2].

4

Notes 1. Sterilize the rice starch by heating it at 150 °C for 2 h. 2. Collected samples should be handled with sterile equipment until the sample is inoculated into the appropriate media. 3. It is advisable to incubate the plates in the normal position to avoid the drop off of the medium bearing the inoculum or to ensure that the suspension does not run down onto the lid of the plate. 4. Inoculate the protist culture in 10–20 tubes in one attempt during axenicization because it may be possible that some protist cultures will not grow, or some will be contaminated.

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References 1. Kalinina LV, Page FC (1992) Culture and preservation of naked amoebae. Acta Protozool 31: 115–126 2. Jensen T, Barnes WG, Meyers D (1970) Axenic cultivation of large populations of Acanthamoeba castellanii (JBM). J Parasitol:904–906

3. Bowers B, Korn ED (1969) The fine structure of Acanthamoeba castellanii (Neff strain) II. Encystment. J Cell Biol 41:786–805

Part V Identification Techniques for Soil Protists

Chapter 21 Classical Wet Mount Method for Observing Live Protists Komal A. Chandarana and Natarajan Amaresan Abstract Protist ecology has used optical microscopes for observing cell features since very beginning. A dissecting microscope with light- and dark-field illumination is ideal for observing protists. The basic method adopted for observing live protists under the microscope is classical wet mount method. Though it is a very primitive method, it is an essential part of the protistological studies till today for observing live cell morphology, locomotion, and its life stage (trophozoites or cyst). Key words Wet mount, Dissecting microscope, Primitive method, Live protists, protistological studies

1

Introduction Wet mount is prepared in order to accurately look at the protistan cell under the dissecting microscope. A wet mount is prepared by placing a fluid having protistan cell and then covering the solution and the specimen with a cover slip. As the pressure of the cover slip increases slowly on the protistan cell, the live cell becomes less mobile and more transparent. Hence, first the movement of the cell can be recorded, followed by location of main cell organelles, such as nucleus and oral apparatus, contractile vacuole, etc., and then micronucleus under the low and high magnification of the microscope [1]. The shape of the cell may be altered by this procedure with the course of the time. Therefore, it is essential to observe the specimen quickly under the low objective to note the shape of the cell. Though investigation with low objective requires expertise in microscopy of protistan cell, it guarantees undamaged cells for records. Video microscopy can also be recorded using this technique with the help of compound microscope equipped with attached camera.

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_21, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Materials Grease-free glass slide, cover slip, Vaseline, dissecting microscope

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Methods 1. Place about 0.5 mL of raw sample on a glass slide, and pick out the desired protistan cell with the help of micropipette or microcapillary while seeing under the 10× objective of dissecting microscope (see Note 1). 2. Transfer this collected protist cell onto a slide which is now with small amount of fluid (see Note 2). 3. Take a coverslip, and place a small dabs of Vaseline to each of four corners of a coverslip. 4. Place this coverslip on a slide containing droplet of protist. 5. Press genteelly four corners of the coverslip with the help of needle, until the protistan cell held firmly between the slide and the coverslip (see Note 3). 6. Place a slide under the microscope, and focus the cell first using low objective.

4

Notes 1. The protistan cell can also be picked directly from the Petri dish under the 10× objective of the microscope. 2. Make sure that droplet containing protistan cell is not too small or too large that fluid comes out of the coverslip. 3. Be kind enough for pressing the coverslip; otherwise the shape of the organism may deteriorate because some of the protist are fragile that cannot withstand the pressure created between the slide and the coverslip.

Reference 1. Lee JJ, Soldo AT (1992) Protocols in protozoology. Society of Protozoologists, Allen

Chapter 22 SEM: Sample Preparation, Fixation, and Staining of Protists Shraddha Saha Abstract The ultrastructure and morphology of the protist can be studied using high-resolution electron microscope, i.e., scanning electron microscope (SEM). One of the greatest advantages of using scanning electron microscope is that observation is possible without any complicated pre-treatment that includes drying sample or coating of sample with metal. Additionally, the findings allow specialists to document details about the 3D structure of protists. With this aim, this chapter describes the procedure pertaining to sample preparation, fixation, and staining of the protists. Key words Morphology, Ultrastructure, Protist, Scanning electron microscope

1

Introduction The scanning electron microscope (SEM) scans the surface through probe of electrons across the surface and generates threedimensional appearance of the specimen. Additionally, SEM provides wide range of magnification of about 15–150,000 with a resolution of order > family > genus > species. 5. Check to see if specimen is represented in the Barcode of Life Database, BOLD (www.boldsystems.org) or GenBank (www. ncbi.nlm.nih.gov). 6. Search by entering genus and species names in the search bar at top right. If the species is represented in the database, the “Taxonomy Browser” will list the number and sources of specimen records. 7. Click on “Download Public Sequences” for a FASTA file of available barcode sequences. 8. Click “Taxonomy Browser” at the top left to explore barcode records by group.

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Notes 1. The forward and reverse primers can be designed from reference sequence available on GenBank. 2. The PCR conditions were as described by Fahrni et al. [29]. 3. The Barcode of Life Data System (BOLD) is an informatics workbench aiding the acquisition, storage, analysis, and publication of DNA barcode records. By assembling molecular, morphological, and distributional data, it bridges a traditional bioinformatics chasm. 4. Users can verify the accuracy of sequence editing by downloading and analyzing these files. HTS platforms do not generate chromatograms which is why only sequence files in text format (FASTA) are uploaded.

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References 1. Margulis L (1993) Symbiosis in cell evolution, 2nd edn. Freeman, NewYork 2. Fenchel T (2002) Origin and early evolution of life. Oxford University Press, Oxford 3. Foster K (2003) Making a robust bimolecular time scale for phylogenetic studies. Protist 154: 43–55 4. Pawlowski J, Audic S, Adl S, Bass D, Belbahri L, Berney C et al (2012) CBOL protist working group: barcoding eukaryotic richness beyond the animal, plant, and fungal kingdoms. PLoS Biol 10:e1001419 5. Countway P, Gast R, Savai P, Caron D (2005) Protistan diversity estimates based on 18s rdna from seawater incubations in the western North Atlantic. J Eukaryot Microbiol 52:95– 106 6. Martı´nez J (2013) Bacterial pathogens: from natural ecosystems to human hosts. Environ Microbiol 15(2):325–333 7. Gao Z, Karlsson I, Geisen S, Kowalchuk G, Jousset A (2019) Protists: puppet masters of the rhizosphere microbiome. Trends Plant Sci 24:165–176 8. Oliverio AM, Geisen S, Delgado-Baquerizo M, Maestre FT, Turner BL, Fierer N (2020) The global-scale distributions of soil protists and their contributions to belowground systems. Sci Adv 6:eaax8787 9. Amacker N, Gao Z, Hu J, Jousset AL, Kowalchuk GA, Geisen S (2022) Protist feeding patterns and growth rate are related to their predatory impacts on soil bacterial communities. FEMS Microbiol Ecol 98:fiac057 10. Mora C, Tittensor D, Adl S, Simpson A, Worm B (2011) How many species are there on Earth and in the ocean? PLoS Biol 9:e1001127 11. Hebert PDN (2004) Identification of birds through DNA barcodes. PLoS Biol 2:312–319 12. Dawson S, Pace N (2002) Novel kingdom level eukaryotic diversity in anoxic environments. Proc Natl Acad Sci U S A 99:8324–8329 13. Lo´pez-Garcı´a P, Rodrı´guez-Valera F, Pedro´sAlio´ C, Moreira D (2001) Unexpected diversity of small eukaryotes in deep-sea Antarctic plankton. Nature 409:603–607 14. Berney C, Fahrni J, Pawlowski J (2004) How many novel eukaryotic “kingdoms”? Pitfalls and limitations of environmental DNA surveys. BMC Biol 2:13 15. Kim E, Harrison J, Sudek S, Jones M, Wilcox H (2011) Newly identified and diverse plastidbearing branch on the eukaryotic tree of life. Proc Natl Acad Sci U S A 108:1496–1500

16. Plassart P, Terrat S, Thomson B, Griffiths R, Dequiedt S, Lelievre M et al (2012) Evaluation of the ISO standard 11063 DNA extraction procedure for assessing soil microbial abundance and community structure. PLoS One 7: e44279 17. Dykova´ I, Boha´cˇova´ L, Fiala I, Macha´cˇkova´ B, Peckova´ H, Dvorˇa´kova´ H (2005) Amoebae of the genera Vannella Bovee, 1965 and Platyamoeba page, 1969 isolated from fish and their phylogeny inferred from SSU rRNA gene and ITS sequences. Eur J Protistol 41:219–230 18. Kosakyan A, Heger TJ, Leander BS, Todorov M, Mitchell EA et al (2012) COI barcoding of nebelid testate amoebae (Amoebozoa: Arcellinida): extensive cryptic diversity and redefinition of the hyalospheniidae schultze. Protist 163(415):434 19. Folmer O, Black M, Hoeh W, Lutz R, Vrijenhoek R (1994) DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol Mar Biol Biotechnol 3:294–299 20. Hagino K, Bendif EM, Young JR, Kogame K, Probert I, Takano Y et al (2011) New evidence for morphological and genetic variation in the cosmopolitan coccolithophore emiliania huxleyi (prymnesiophyceae) from the COX1bATP4 genes 1. J Phycol 47:1164–1176 21. Virginia M, Puerta S, Bachvaroff T, Delwiche C (2004) The complete mitochondrial genome sequence of the haptophyte Emiliania huxleyi and its relation to heterokonts. DNA Res 11:1– 10 22. Barth D, Krenek S, Fokin SI, Berendonk TU (2006) Intraspecific genetic variation in Parame cium revealed by mitochondrial cytochrome C oxidase I sequences. J Eukaryot Microbiol 53:20–25 23. Chantangsi C, Lynn DH, Brandl MT, Cole JC, Hetrick N et al (2007) Barcoding ciliates: a comprehensive study of 75 isolates of the genus Tetrahymena. Int J Syst Evol Microbiol 57:2412–2425 24. Jerome CA, Lynn DH (1996) Identifying and distinguishing sibling species in the Tetrahymena pyriformis complex (Ciliophora, Oligohymenophorea) using PCR/RFLP analysis of nuclear ribosomal DNA. J Eukaryot Microbiol 43:492–497 25. Medlin L, Elwood HJ, Stickel S, Sogin ML (1988) The characterization of enzymatically amplified eukaryotic 16S-like rRNA-coding regions. Gene 71:491–499

DNA Barcoding Techniques for Protists 26. Hebert P, Braukmann T, Prosser S (2018) A sequel to Sanger: amplicon sequencing that scales. BMC Genomics 19:219 27. Saunders GW, McDevit DC (2012) Methods for DNA barcoding photosynthetic protists emphasizing the macroalgae and diatoms. In: Kress W, Erickson D (eds) DNA barcodes, Methods in molecular biology, vol 858. Humana Press, Totowa, pp 207–222

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28. www.boldsystems.org 29. Fahrni JH, Bolivar I, Berney C, Nassonova E, Smirnov A, Pawlowski J (2003) Phylogeny of lobose amoebae based on actin and smallsubunit ribosomal RNA genes. Mol Biol Evol 20:1881–1886

Chapter 30 Unequivocal Identification Through Fluorescence In Situ Hybridization (FISH) Technique Jean C. V. Dutra and Maria C. P. Batitucci Abstract Protists represent the largest portion of the diversity of eukaryotes in the globe and have been recognized for performing indispensable activities in biogeochemical cycles. With the advancement of molecular biology techniques, this group of living beings has been widely studied. Among molecular techniques, FISH assay stands out, which, through the hybridization of specific probes, provides an unequivocal phylogenetic relationship. Hence here we present the FISH assay, indicating step by step how to successfully perform the in situ hybridization process. Key words Soil protist, FISH assay, Fluorescence hybridization, In situ hybridization

1

Introduction Faced with environmental changes across the globe, studies on biodiversity have grown, especially studies on protists. Despite being relatively small organisms, protists have been recognized for performing indispensable activities in biogeochemical cycles, being crucial in most ecosystems [1]. These microorganisms represent the largest portion of the diversity of eukaryotes and inhabit all environments on the planet [2–5]. So, understanding the diversity of protists and their relationship with other organisms, integrating different ecosystems, is essential to have a holistic view of ecosystems [1]. The diversity of protists and the structure of communities vary according to the habitat, especially those that inhabit the soil, as they act as bioindicators and provide information about environmental conditions [6, 7]. In recent years, the great diversity of soil protists has ceased to be underestimated due to advances in molecular biology techniques [8, 9].

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_30, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Among the molecular techniques used to assess the diversity of protists, the fluorescent in situ hybridization assay (FISH assay) stands out. FISH technique is considered a powerful tool by complementary methods of more traditional DNA detection, such as the methods based on PCR. Through the FISH assay, it is possible to reveal the tissue location of an infection and the host response, as in traditional histopathological assays, maintaining a high degree of specificity and sensitivity, similar to the observed in the PCR technique. In this way, successful hybridization can provide unequivocal phylogenetic confirmation that a specific pathogen is associated with a specific host tissue [10]. Regarding molecular techniques for identification of organisms, in this chapter, we will explain the unequivocal method of soil protist identification by the FISH assay.

2

Materials In order to avoid contamination, it is recommended to use sterile material and solutions from reliable sources.

2.1 Basic Requirements 2.2

Solutions

Sterile Petri dish, micropipette and tips, Pasteur pipette.

1. Bouin’s solution. 2. Ethanol. 3. Hybridization buffer (see Note1). 4. Probes. 5. Anti-fade mounting medium. 6. DAPI.

2.3

3

Equipment

Epifluorescence microscope.

Methods

3.1 Preparation of Protist Cells

The organisms (specimens of protists) studied can be isolated using appropriate techniques or can be provided by partner laboratories. 1. The protist organisms to be studied must be transferred, with the aid of a micropipette, to a sterile Petri dish containing sterile water (see Note 2). 2. Using a sterile Pasteur pipette, transfer the cells to sterile tubes with a lid, gently agitate the cells with the micropipette, centrifuge at 860 rcf for 10 min, and discard the supernatant. Add sterile water and repeat this process two to five times (see Note 3).

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3. Fix the cells with Bouin’s solution at 50% final concentration, and, to perform the fluorescence in situ hybridization (FISH) assay, transfer up to 20 individuals to sterile slides, and allow to air dry at room temperature [11]. 3.2

FISH Assay

The method described below is the whole-cell hybridization as described by Gong et al. [11], in which protist cells are subjected to hybridization with a specific probe and revealed by fluorescence. 1. Following the last process described, wash the slides with distilled water. 2. Progressively dehydrate the cells by washing the slides with an ethanol gradient (30%, 50%, 80%, and 100%). 3. Incubate the samples on slides at 46 °C for 3 h in hybridization buffer (buffer plus specific probes at a concentration of 5 ng μL-1) (see Notes 4 and 5). 4. At the end of the hybridization, wash the slides with a buffer for 15 min at 48 °C, and then rinse with double-distilled water. 5. Mounting of slides proceeds with the addition of anti-fade mounting medium mixed with DAPI (50 ng mL-1). 6. After mounting the slide, observe under an epifluorescence microscope, with UV excitation for DAPI signals (see Note 6). 7. Take photographs with the help of an attached camera.

4

Notes 1. The hybridization buffer is composed of 20 mM Tris-HCl at pH 8.0, 0.9 M NaCl, 0.01% SDS (sodium dodecyl sulfate) and 30% formamide. Prepare the hybridization buffer as described by Omar et al. [11]. 2. For marine organisms use autoclaved seawater, and for freshwater organisms use twice-distilled freshwater. 3. Cell washing is recommended to remove microorganisms adhered to the cell surface of protists and minimize contamination. 4. A mix of probes specific to the analyzed material must be used to ensure hybridization and assay success. In this way, in addition to the target probes, nonsense probes can be used as a negative control. 5. MathFISH (web-based tool) can be used to assess the sensitivity and specificity of the probes, as well as estimating the optimal formamide concentration (40%) to optimize mismatch discrimination [12]. 6. Photomicrographs can be taken at this step, and data analysis must be done using suitable software.

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Acknowledgments Authors are grateful to Ufes (Universidade Federal do Espı´rito Santo) and to FAPES (Fundac¸˜ao de Amparo a` Pesquisa e Inovac¸˜ao do Espı´rito Santo). References 1. Burki F, Sandin MM, Jamy M (2021) Diversity and ecology of protists revealed by metabarcoding. Curr Biol 31:R1267–R1280 2. Keeling PJ, Burki F (2019) Progress towards the tree of eukaryotes. Curr Biol 29:R808– R817 3. Geisen S, Mitchell EAD, Adl S et al (2018) Soil protists: a fertile frontier in soil biology research. FEMS Microbiol Rev 42:293–323 4. Bar-On YM, Phillips R, Milo R (2018) The biomass distribution on Earth. Proc Natl Acad Sci 115:6506–6511 5. Caron DA, Countway PD, Jones AC et al (2012) Marine protistan diversity. Annu Rev Mar Sci 4:467–493 6. Foissner W (1997) Protozoa as bioindicators in agroecosystems, with emphasis on farming practices, biocides, and biodiversity. Agric Ecosyst Environ 62:93–103 7. Payne RJ (2013) Seven reasons why protists make useful bioindicators. Acta Protozool 52: 115–128

8. Foissner W (1999) Protist diversity: estimates of the near-imponderable. Protist 150:363– 368 9. Mahe´ F, de Vargas C, Bass D et al (2017) Parasites dominate hyperdiverse soil protist communities in Neotropical rainforests. Nat Ecol Evol 1:0091 10. Carnegie RB, Barber BJ, Distel DL (2003) Detection of the oyster parasite Bonamia ostreae by fluorescent in situ hybridization. Dis Aquat Org 55:247–252 11. Omar A, Zhang Q, Zou S et al (2017) Morphology and phylogeny of the soil ciliate Metopus yantaiensis n. sp.(Ciliophora, Metopida), with identification of the intracellular bacteria. J Eukaryot Microbiol 64:792–805 12. Yilmaz LS, Parnerkar S, Noguera DR (2011) mathFISH, a web tool that uses thermodynamics-based mathematical models for in silico evaluation of oligonucleotide probes for fluorescence in situ hybridization. Appl Environ Microbiol 77:1118–1122

Part VI Other Protistan

Chapter 31 Isolation, DNA Extraction, Amplification, and Gel Electrophoresis of Single-Celled Nonmarine Foraminifera (Rhizaria) Maria Holzmann Abstract Molecular tools are an important part to study the phylogeny and taxonomy of foraminifera. In the present chapter, isolation and extraction of these single-celled organisms will be described. Furthermore, the amplification of the 18S barcoding fragment in foraminifera and detection of amplified products by agarose gel electrophoresis will be detailed. Key words Nonmarine foraminifera, DNA extraction, PCR amplification, Gel electrophoresis

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Introduction Foraminifera are a group of protists belonging to Rhizaria. They are a primarily marine taxon but are known from freshwater and terrestrial environments since the nineteenth century when European and North American protistologists described the first species [1–3]. With the advent of molecular systematics, several new species, genera, and families have been described based on morphological and molecular methods [4–9]. Additionally, metabarcoding studies have revealed a large diversity of foraminifera in environmental samples taken from freshwater and soil [10–12]. A metatranscriptomic study confirmed the widespread occurrence of foraminifera in soils [13]. Freshwater and terrestrial foraminifera are single-chambered (monothalamid) protists and include three basic morphological types: with a flexible, organic-walled test, with a finely agglutinated test, and without a test (athalamid). They possess only few morphological features that can be used for distinction making molecular identification essential for their correct classification [7]. Prior

N. Amaresan and Komal A. Chandarana (eds.), Practical Handbook on Soil Protists, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-3750-0_31, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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forward primers A10

6F

14F3 14F1 15A 1000Bp

reverse primers

7R s12r

SSU

2000Bp s17r

3000Bp 20r sB

Fig. 1 Diagram of the 18S gene in foraminifera with approximate positions of amplification primers

to sequencing foraminifera from freshwater or soil samples, individuals have to be isolated, and their DNA has to be extracted and amplified by PCR. It is essential to choose living foraminifera that are distinguished by pseudopodial activity [14]. In foraminifera, the granuloreticulopodial network is used for movement, adhesion to the substrate, feeding, and test formation and extends from one or several apertures. The streaming of the pseudopodial network can be observed under a binocular or a microscope, depending on the size of the cells. Foraminifera feed on a wide variety of organisms including bacteria, flagellates, phototrophic algae, heterotrophic protists, and yeast [8]. Consequently, most foraminiferal DNA extractions will contain extraneous DNA originating from partially digested food organisms or epibiontic microorganisms living on the surface of foraminiferal tests. Specific foraminiferal primers have therefore to be used to ensure that the amplified genomic fragment originated from foraminifera and not from other microorganisms [14]. The barcoding fragment of foraminifera consists of the 3′end part of the SSU rDNA gene and comprises six variable segments (Fig. 1). Three of them are specific to foraminifera, while the three others correspond to typical eukaryotic variable regions [15]. Foraminiferal barcodes are used for species identification and to investigate phylogenetic relationships. More than 9000 18S sequences of foraminifera are publicly available at NCBI GenBank (https:// www.ncbi.nlm.nih.gov/). Mini-barcodes for foraminifera have also been developed for environmental monitoring using nextgeneration sequencing technologies [16].

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Materials Wear laboratory gloves for each step. Use ultrapure water when preparing solutions for DNA extraction and amplification. Prepare and store all reagents at room temperature unless otherwise indicated. Use sterile filter tips. Follow waste disposal regulations when disposing of waste material.

Isolation, DNA Extraction, Amplification, and Gel Electrophoresis. . .

2.1

Isolation

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1. Sterile Petri dishes (9 cm diameter). 2. Glass beaker 50 mL filled with deionized water (wash bottle with 70% ethanol). 3. Natural hairbrushes in different sizes (3/0, 5/0, 6/0). 4. Pipette and sterile pipette tips with filter (20 uL). 5. Binocular or microscope, preferentially equipped with a camera.

2.2 Guanidine Extraction

1. Extraction tubes (0.5 mL) and pestles; lab storage rack; isopropanol; GlycoBlue; 70% ethanol; TE buffer; pH 8.0 (molecular grade); sterile filter tips for pipetting; heating block (60 °C, 30 °C); centrifuge: maximum speed 14,000 rpm; freezer (-20 °C). 2. Guanidine Solution for a Total Volume of 212 mL. 100 mL ddH2O. 100 g guanidinium isothiocyanate. 10.6 mL Tris 1 M pH 7.6 4.25 mL EDTA 0.5 M Add a stir bar to the solution, and put it on a magnetic stirrer, warm 10 min at 70 °C, dissolve by stirring then add: 4.24 g of Sarkosyl (N-Lauroylsarcosine Sodium Salt) 2.1 mL β-Mercaptoethanol Adjust to 212 mL with distilled H2O and preserve at 4 °C.

2.3

Amplification

1. PCR tubes (0.2 mL), 0.5 mL or 1.5 mL tubes (depending on the amount of PCR master mix), pipettes and sterile filter tips, vortex mixer, cooling rack or ice bucket, thermocycler. 2. PCR reagents: ddH2O, 10X buffer, dNTP’s solution, primers 10 pM (2X), BSA, Taq polymerase, DNA extracts.

2.4 Agarose Gel Electrophoresis

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1. Power supply, horizontal gel tray, horizontal gel tank, gel combs, UV light source, microwave, glass beaker, agarose gel (1.5%): 4.5 g agarose +30 mL TAE (1X), TAE buffer (1X), SYBR Safe DNA gel stain (Invitrogen), loading dye, molecular weight marker.

Methods

3.1 Isolation of Foraminifera from Samples

1. Transfer freshwater sediment samples into Petri dishes and cover them with water from the sampling site. Soil samples are also transferred to Petri dishes and covered with natural mineral water (f.i. Volvic). The bottom of the Petri dishes should only be lightly covered with sediment. Most foraminiferal species cannot be cultured. Some attempts, however, have been made with freshwater foraminifera that could be

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maintained under laboratory conditions for a few weeks [9]. Observe foraminifera using a binocular or microscope, and select living specimens characterized by pseudopodial activity. Living foraminifera are often surrounded by debris; try to carefully clean specimens with a brush prior to extraction (see Note 1). As foraminiferal cells are destroyed in the extraction process, it is preferable to take images/videos and do measurements prior to extraction. 2. For extraction, take up a foraminiferal cell with the tip of the brush, and transfer it into the guanidine solution (see Note 2). Before picking up the next cell, rinse the brush with 70% ethanol and then with deionized water to minimize contamination. 3. If cells are very small (