Staphylococcus aureus: Methods and Protocols (Methods in Molecular Biology, 2341) 1071615491, 9781071615492

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Methods in Molecular Biology 2341

Kelly C. Rice Editor

Staphylococcus aureus Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Staphylococcus aureus Methods and Protocols

Edited by

Kelly C. Rice IFAS, Department of Microbiology and Cell Science, University of Florida, Gainesville, FL, USA

Editor Kelly C. Rice IFAS, Department of Microbiology and Cell Science University of Florida Gainesville, FL, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1549-2 ISBN 978-1-0716-1550-8 (eBook) https://doi.org/10.1007/978-1-0716-1550-8 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved 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.

Preface The field of staphylococcal biology is now entrenched within the post-genomic era, in which high-throughput sequencing of genomes and transcriptomes has become both technically and financially accessible to many researchers. These powerful technologies have generated massive amounts of genomic and global transcriptome (RNAseq) data, which, when paired with proteomics and metabolomics analyses of S. aureus, covers all aspects of the central dogma relevant to this pathogen’s physiology and virulence potential. In parallel to these omics methods, our genetic repertoire for studying S. aureus has been significantly expanded in recent years by readily available screening resources such as the Nebraska Transposon Mutant Library (NTML; containing nearly 2000 defined transposon mutations in nonessential genes) and its corresponding series of allele-exchange plasmid tools for transposon replacement with alternative antibiotic resistance or fluorescent reporter genes. We have also seen impressive leaps forward in tools for conducting transposon sequencing (TnSeq) to directly assess the importance of S. aureus genes for fitness in vivo. Likewise, techniques such as dual RNAseq have allowed direct assessment of interaction of S. aureus with host during in vivo infection. Increasingly, these studies have found that, in addition to traditional virulence traits commonly associated with S. aureus pathogenesis (cell surface adhesins, secreted toxins/tissue-degrading enzymes, biofilm formation), mutations affecting bacterial metabolism have a profound influence on the ability of this organism to survive in vivo. This explosion in omics and high-throughput data has also facilitated the follow-up discovery and characterization of genes of unknown function (the dreaded “hypothetical protein” genome annotations!) at an unprecedented speed. In mining these big data sets for genes important to S. aureus physiology and virulence, a typical next step in a laboratory’s research pipeline is to create a clean isogenic mutant and complement strain in a gene of interest, followed by genetic and in vitro/in vivo phenotypic validation and characterization of mutant phenotypes. The molecular biology methodologies required for S. aureus mutant making and genetic analyses have already been well covered in Methods in Molecular Biology Book volume 1373: The Genetic Manipulation of Staphylococci. Therefore, the overall goal of this current volume is to provide a companion technical resource for phenotypic characterization of S. aureus mutants, focusing on in vitro and ex vivo methodologies. To this end, in vitro assessment of classical S. aureus virulence attributes (pigmentation, cell clumping, and assessment of secreted hemolysin, nuclease, murein hydrolase, and protease activities) is presented in the first five chapters. These are followed by a chapter focused on quantifying promoter activity using a S. aureus codon-optimized lacZ plasmid, a useful tool for assessing the effect of putative regulators on expression of genes of interest. Chapters 7–13 cover key in vitro phenotypic assays that measure aspects of S. aureus physiology (membrane fluidity, membrane potential, O2 consumption, antibiotic tolerance, metabolomics) and biologically relevant growth environments (growth as biofilm and under low shear) that have potential to impact the virulence and stress resistance of this pathogen. The final five chapters are dedicated to in vivo (harvesting samples from a biofilm infection

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model for S. aureus proteomics) and ex vivo (S. aureus survival within primary macrophages and in whole blood, killing of host cells, and colonization of human skin) models of hostpathogen interaction. Gainesville, FL, USA

Kelly C. Rice

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

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1 Measurement of Staphylococcus aureus Pigmentation by Methanol Extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Leia E. Sullivan and Kelly C. Rice 2 Analysis of Murein Hydrolases and Proteases Through Zymography . . . . . . . . . . 9 Nichole A. Seawell and Jeffrey L. Bose 3 Detection and Quantification of Secreted Nuclease Activity in Staphylococcus aureus Culture Supernatants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 Richard E. Wiemels, Rebecca A. Keogh, and Ronan K. Carroll 4 Quantitative Hemolysis Assays. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 Miranda J. Ridder, Seth M. Daly, Pamela R. Hall, and Jeffrey L. Bose 5 In Vitro Assay for Quantifying Clumping of Staphylococcus aureus . . . . . . . . . . . . 31 Heidi A. Crosby, Jakub M. Kwiecinski, and Alexander R. Horswill 6 Measuring Staphylococcal Promoter Activities Using a Codon-Optimized β-Galactosidase Reporter. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37 Christina N. Krute, Nichole A. Seawell, and Jeffrey L. Bose 7 Evaluation of Staphylococcus aureus Antibiotic Tolerance Using Kill Curve Assays. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Jessica N. Brandwein and Kelly C. Rice 8 Fluorescence Polarization (FP) Assay for Measuring Staphylococcus aureus Membrane Fluidity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Kiran B. Tiwari, Suranjana Sen, Craig Gatto, and Brian J. Wilkinson 9 Quantification of Staphylococcus aureus Biofilm Formation by Crystal Violet and Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Adam B. Grossman, Dylan J. Burgin, and Kelly C. Rice 10 Growth of Staphylococcus aureus Using a Rotary Cell Culture System. . . . . . . . . . 79 Matthew R. Hauserman and Kelly C. Rice 11 Time-Resolved Fluorescence Assay for Measuring Oxygen Consumption Rates in Staphylococcus aureus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Logan L. Bulock, Jennifer L. Endres, and Marat R. Sadykov 12 Quantifying Staphylococcus aureus Membrane Potential Using Flow Cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Neal D. Hammer 13 Best Practices for Preparation of Staphylococcus aureus Metabolomics Samples. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Kimberly L. James and Kelly C. Rice 14 A Rat Model of Orthopedic Implant-Associated Infection for Identification of Staphylococcal Biofilm Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Mei G. Lei, Ravi Kr. Gupta, and Chia Y. Lee

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Contents

An Ex Vivo Model for Assessing Growth and Survivability of Staphylococcus aureus in Whole Human Blood. . . . . . . . . . . . . . . . . . . . . . . . . . . . Brittney D. Gimza, Stephanie M. Marroquin, and Lindsey N. Shaw Bone Marrow-Derived Macrophage Infection Assay. . . . . . . . . . . . . . . . . . . . . . . . . Miranda J. Ridder, Mary A. Markiewicz, and Jeffrey L. Bose Measurement of Osteoblast Cytotoxicity Induced by S. aureus Secreted Toxins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caleb A. Ford and James E. Cassat Human Skin In Vitro Colonization Model for a Skin Wound Infected by Staphylococcus aureus Biofilm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jenelle E. Chapman and Michael E. Olson

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors JEFFREY L. BOSE • Department of Microbiology, Molecular Genetics and Immunology, The University of Kansas Medical Center, Kansas City, KS, USA JESSICA N. BRANDWEIN • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA LOGAN L. BULOCK • Department of Pathology and Microbiology, Center for Staphylococcal Research, University of Nebraska Medical Center, Omaha, NE, USA DYLAN J. BURGIN • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA RONAN K. CARROLL • Department of Biological Sciences, Ohio University, Athens, OH, USA JAMES E. CASSAT • Department of Biomedical Engineering, Vanderbilt University, Nashville, TN, USA; Division of Pediatric Infectious Diseases, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, TN, USA; Department of Pathology, Microbiology, and Immunology, Vanderbilt University Medical Center, Nashville, TN, USA; Vanderbilt Institute for Infection, Immunology and Inflammation, Vanderbilt University Medical Center, Nashville, TN, USA; Vanderbilt Center for Bone Biology, Vanderbilt University Medical Center, Nashville, TN, USA JENELLE E. CHAPMAN • Department of Medical Microbiology, Immunology and Cell Biology, Southern Illinois University School of Medicine, Springfield, IL, USA HEIDI A. CROSBY • Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA; New England Biolabs Inc, Ipswich, MA, USA SETH M. DALY • Department of Pharmaceutical Sciences, University of New Mexico College of Pharmacy, Albuquerque, NM, USA JENNIFER L. ENDRES • Department of Pathology and Microbiology, Center for Staphylococcal Research, University of Nebraska Medical Center, Omaha, NE, USA CALEB A. FORD • Department of Biomedical Engineering, Vanderbilt University, Nashville, TN, USA CRAIG GATTO • School of Biological Sciences, Illinois State University, Normal, IL, USA BRITTNEY D. GIMZA • Department of Cell Biology, Microbiology, and Molecular Biology, University of South Florida, Tampa, FL, USA ADAM B. GROSSMAN • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA RAVI KR. GUPTA • Department of Microbiology and Immunology, University of Arkansas for Medical Sciences, Little Rock, AR, USA; Department of Environmental Microbiology, Babasaheb Bhimrao Ambedkar University, Lucknow, India PAMELA R. HALL • Department of Pharmaceutical Sciences, University of New Mexico College of Pharmacy, Albuquerque, NM, USA NEAL D. HAMMER • Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI, USA MATTHEW R. HAUSERMAN • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA ALEXANDER R. HORSWILL • Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA; Department of Veterans Affairs, Eastern Colorado Health Care System, Denver, CO, USA

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KIMBERLY L. JAMES • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA REBECCA A. KEOGH • Department of Biological Sciences, Ohio University, Athens, OH, USA CHRISTINA N. KRUTE • Department of Microbiology, Molecular Genetics and Immunology, The University of Kansas Medical Center, Kansas City, KS, USA JAKUB M. KWIECINSKI • Department of Immunology and Microbiology, University of Colorado School of Medicine, Aurora, CO, USA CHIA Y. LEE • Department of Microbiology and Immunology, University of Arkansas for Medical Sciences, Little Rock, AR, USA MEI G. LEI • Department of Microbiology and Immunology, University of Arkansas for Medical Sciences, Little Rock, AR, USA MARY A. MARKIEWICZ • Department of Microbiology, Molecular Genetics and Immunology, The University of Kansas Medical Center, Kansas City, KS, USA STEPHANIE M. MARROQUIN • Department of Cell Biology, Microbiology, and Molecular Biology, University of South Florida, Tampa, FL, USA MICHAEL E. OLSON • Department of Medical Microbiology, Immunology and Cell Biology, Southern Illinois University School of Medicine, Springfield, IL, USA KELLY C. RICE • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA MIRANDA J. RIDDER • Department of Microbiology, Molecular Genetics and Immunology, The University of Kansas Medical Center, Kansas City, KS, USA MARAT R. SADYKOV • Department of Pathology and Microbiology, Center for Staphylococcal Research, University of Nebraska Medical Center, Omaha, NE, USA NICHOLE A. SEAWELL • Department of Microbiology, Molecular Genetics and Immunology, The University of Kansas Medical Center, Kansas City, KS, USA SURANJANA SEN • Loyola University Chicago, Chicago, IL, USA LINDSEY N. SHAW • Department of Cell Biology, Microbiology, and Molecular Biology, University of South Florida, Tampa, FL, USA LEIA E. SULLIVAN • Department of Microbiology and Cell Science, IFAS, University of Florida, Gainesville, FL, USA KIRAN B. TIWARI • School of Biological Sciences, Illinois State University, Normal, IL, USA RICHARD E. WIEMELS • Department of Biological Sciences, Ohio University, Athens, OH, USA BRIAN J. WILKINSON • School of Biological Sciences, Illinois State University, Normal, IL, USA

Chapter 1 Measurement of Staphylococcus aureus Pigmentation by Methanol Extraction Leia E. Sullivan and Kelly C. Rice Abstract Many S. aureus strains produce membrane-associated carotenoid pigments, advantageous secondary metabolites that can alter membrane fluidity, resistance to antimicrobial peptides (AMPs) and act as antioxidants, properties that can impact resistance against aspects of the host innate immune system. Several studies have reported connections between mutations in both regulatory (i.e., alternative sigma factor B) and metabolic (purine biosynthesis, oxidative phosphorylation) genes, and noticeable differences in carotenoid pigmentation. This chapter outlines a simple protocol to quantify cellular pigments using a methanol extraction method. Key words Staphylococcus aureus, Carotenoids, Membrane fluidity, Pigment, Oxidative stress resistance

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Introduction Pigment producing bacteria are ubiquitously found in nature. Staphylococcus aureus, so-named by Dr. Rosenbach for its golden (“aurum”) colonies on agar plates [1], produces 17 different triterpenoid carotenoids [2]. These carotenoids are unique in that they contain a C30 chain with alternating single and double bonds, contrary to the C40 chains found in most other organisms [2]. Most recognized as contributing to the orange/gold pigment of this bacterium is the membrane-associated carotenoid staphyloxanthin, whose biosynthesis is encoded by the crtOPQMN operon [3, 4]. Staphyloxanthin (α-D-glucopyranosyl 1-O-(4,40 -diaponeurosporen-4-oate)-6-O-(12-methyltetradecanoate)) is produced in a 5-step enzymatic reaction that begins with condensation of two farnesyl diphosphate molecules (catalyzed by CrtM, a dehydrosqualene synthase), followed by two stepwise oxidation reactions catalyzed by CrtN and CrtP, and two glucose esterification steps at the C100 and C600 positions (catalyzed by CrtQ and CrtO, respectively) [4].

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Leia E. Sullivan and Kelly C. Rice

Expression of the crt operon is positively regulated by alternative sigma B [4], MsaB [5], the AirSR two-component system [6], and the predicted thioredoxin-like oxidoreductase YjbH [7, 8], whereas crt expression is repressed by the small RNA SsrA [9]. Additionally, pigmentation of S. aureus is affected both positively and negatively by the perturbation of several metabolic pathways, including mutation of genes involved in purine biosynthesis [10], methionine sulfoxide reductase [11], nitric oxide synthase [12], and various aspects of oxidative phosphorylation [10, 13]. In addition to acting as a phenotypic marker for disruption of the genetic systems and metabolic pathways mentioned above, staphyloxanthin has been shown to be an important mediator of oxidative stress resistance [14, 15]. This protective effect is a function of the numerous conjugated C¼C bonds in the carotenoid backbone, which confers both its pigmentation and antioxidant properties, the latter of which occurs by interaction of the C¼C bonds with various free radicals [16]. An increase in susceptibility to hydrogen peroxide, superoxide, hydroxyl radicals, and neutrophil killing was observed in S. aureus crtM mutants [14, 15]. Additionally, staphyloxanthin content in the S .aureus cell membrane increases membrane rigidity [17, 18], which has been shown to correlate with protection from the killing action of antimicrobial peptides [17]. Pigmentless and low-pigment producing S. aureus strains are also more sensitive to killing by exogenous fatty acids such as oleic acid [19, 20], but production of staphyloxanthin does not appear to impact the fatty acid composition of the staphylococcal membrane [18]. More recently, staphyloxanthin has also been shown to be protective against the killing effects of Cold Atmospheric Plasma treatment [21] and UV radiation [22]. Because of the protective effects of carotenoids against various environmental and host stresses, compounds that inhibit specific aspects of its biosynthetic pathway have been investigated as potential therapeutic agents [23–26]. Likewise, there is evidence that staphyloxanthin provides a protective advantage to S. aureus during infection in vivo. A crtM mutant displayed reduced survival relative to the wild-type strain in a murine skin lesion infection model [15], and S. aureus was more susceptible to innate immune clearance when mouse infections were treated with a CrtN inhibitor [24]. As evidenced from the studies discussed above, pigmentation is related to multiple facets of S. aureus physiology, stress resistance, and virulence. Thus, in many experimental contexts, quantification of this phenotype is desirable, as visual/qualitative assessment on an agar plate streak can be confounded by variations in the inoculum and color of the growth medium. In this chapter, we present a simple, fast, yet sensitive method to quantify S. aureus pigmentation by methanol extraction, followed by

S. aureus Pigment Quantification

3

absorbance measurement, based on the protocol originally published by Ohta and colleagues [27], with adaptations described in [12].

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Materials 1. Tryptic Soy Agar (TSA) plates: 15 g tryptic soy broth (TSB) and 7.5 g agar in 500 ml dH2O, autoclaved for 20–25 min on liquid cycle (121  C) to sterilize. Cool sterilized medium in a water bath to 50–55  C, then pour aliquots into sterile plastic petri dishes. 2. Methanol, molecular grade. 3. dH2O, autoclaved for 20–25 min on liquid cycle (121  C) to sterilize. 4. S. aureus strains to be tested (20% (v/v) glycerol stocks stored at 80  C, or lyophilized stocks). 5. Quartz or plastic disposable cuvettes (1.0 cm path length, 1.5 ml sample volume capacity), compatible with reading wavelengths between 340 and 750 nm. 6. Sterile 1.8 ml microcentrifuge tubes. 7. Sterile toothpicks or sterile pipette tips (200 μl or 1000 μl size). 8. Static incubator set at 37  C. 9. Spectrophotometer with sample holder for 1.5 ml cuvettes and capable of measuring absorbance between 340 and 750 nm. 10. Benchtop microcentrifuge. 11. Waterbath or heat block set to 55  C.

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Methods 1. Streak S. aureus strains of interest onto a TSA plate from stocks. Grow for 48 h at 37  C (see Note 1). 2. Scrape up cells from each TSA plate using a sterile toothpick or sterile pipette tip and resuspend in 1 ml sterile water in a microcentrifuge tube, being careful not to transfer chunk of agar. Scrape up enough cells to get a thick pellet upon centrifugation (approximately ½–¾ of the dense/undiluted bacterial lawn) (see Note 2). 3. Centrifuge all sample tubes at 17,000  g for 3 min at room temperature (25  C). 4. Remove the supernatant from each cell pellet by pipetting and discard liquid in an appropriate biohazard waste receptacle. 5. Resuspend each cell pellet in 1 ml of sterile water.

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6. Centrifuge at 17,000  g for 3 min at room temperature (25  C). 7. Remove the supernatant by pipetting and discard in appropriate biohazard waste receptacle. 8. Resuspend each cell pellet in 420 μl methanol. Scrape and gently pipette up and down until cell pellet is completely disrupted and resuspended (see Note 3). 9. Vortex each tube at top speed for 10 s. Then immediately withdraw 20 μl from each tube and add each to 980 μl H2O in a cuvette. Mix well and read/record OD600 of each sample (see Table 1 for an example of how to record the data). These values reflect the cell density of each sample and will be used to standardize all pigment quantification data as described in Table 1. 10. Incubate remaining 400 μl cell suspension (from step 8) for 5 min at 55  C. This heating step promotes release and solubilization of cellular pigments into the methanol. 11. Centrifuge all tubes for 2 min at 17,000  g at room temperature (25  C). 12. Withdraw 350 μl from each tube and transfer each to a cuvette containing 650 μl methanol. 13. Mix each cuvette well and record absorbance of each sample at 465 nm, which reflects the amount of staphyloxanthin present in each sample (see Note 4). 14. Follow the steps outlined in Table 1 to standardize and calculate the relative differences in pigmentation between samples.

Table 1 Standardization of pigment quantification between samples

Sample

OD600 (step 9) OD600  50 (50-fold (Adjusted dilution) OD600)

Wildtype 0.05 (control) Mutant strain

0.1

Relative OD600 (Adjusted OD600 of sample/ A465 A465/ Adjusted OD600 (step Relative OD600 of control sample) 13)

2.5

1.0

0.5

0.5

5.0

2.0

0.25

0.125

S. aureus Pigment Quantification

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Notes 1. In our experience, pigment quantification is maximal when S. aureus strains are grown for 48 h on TSA plates. However, this protocol can be adapted for other culture conditions as dictated by the environmental parameter, genetic mutation, or other variable(s) under study. We have quantified S. aureus cell pigmentation with this method using TSA plate growth for 24 and 48 h, and in broth cultures grown for 24 and 48 h. In these assays, we have noticed increased pigmentation is measured at both time points when S. aureus is grown on agar plates compared to broth cultures. We have also noticed that S. aureus TSA plate growth in low-oxygen or anaerobic conditions tends to result in less pigmentation compared to growth in aerobic conditions (unpublished observations). 2. To scrape up cells from the TSA plate, we find using a sterile 1 ml pipette tip can remove a lot of cells from the agar plate without disrupting the agar itself. It can be held between two fingers at the wider end and, holding it almost parallel to the agar surface, cells can be gently scraped from the agar surface and then easily dislodged into the microcentrifuge tube of water using a gentle swirling/shaking motion. This step can be repeated several times until enough cells are transferred from plate to tube. 3. At this resuspension step, the cell pellet does not easily disperse/resuspend into methanol. Therefore, it may take several minutes per sample to completely resuspend the cell pellet, which is best accomplished by alteration of gentle scrapping and resuspending with a pipette, and vigorous vortexing. It is important to break up and resuspend the cell pellet as best possible, to enable robust extraction of pigments when sample is heated to 55  C. Keep this step of the protocol consistent for all samples. For example, if 5 min is spent resuspending one cell pellet, subject all of the other cell pellets to this same treatment. 4. Staphyloxanthin in methanol displays peak absorbance at 465 nm [2]. If detection of other pigments is desired, it is recommended to perform a full spectral scan (~300–500 nm) on each sample [2].

References 1. Rosenbach FJ (1884) Mikro-organismen bei den Wund-Infections-Krankheiten des Menschen. J.F. Bergmann, Wiesbaden 2. Marshall JH, Wilmoth GJ (1981) Pigments of Staphylococcus aureus, a series of triterpenoid carotenoids. J Bacteriol 147(3):900–913.

https://doi.org/10.1128/jb.147.3.900-913. 1981 3. Wieland B, Feil C, Gloria-Maercker E et al (1994) Genetic and biochemical analyses of the biosynthesis of the yellow carotenoid 4,40 -diaponeurosporene of Staphylococcus aureus. J Bacteriol 176(24):7719–7726.

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https://doi.org/10.1128/jb.176.24.77197726.1994 4. Pelz A, Wieland KP, Putzbach K et al (2005) Structure and biosynthesis of staphyloxanthin from Staphylococcus aureus. J Biol Chem 280 (37):32493–32498. https://doi.org/10.1074 /jbc.M505070200 5. Pandey S, Sahukhal GS, Elasri MO (2019) The msaABCR operon regulates the response to oxidative stress in Staphylococcus aureus. J Bacteriol 201(21):e00417-19. https://doi.org/ 10.1128/jb.00417-19 6. Hall JW, Yang J, Guo H et al (2017) The Staphylococcus aureus airSR two-component system mediates reactive oxygen species resistance via transcriptional regulation of staphyloxanthin production. Infect Immun 85(2): e00838-16. https://doi.org/10.1128/iai. 00838-16 7. Austin CM, Garabaglu S, Krute CN et al (2019) Contribution of YjbIH to virulence factor expression and host colonization in Staphylococcus aureus. Infect Immun 87(6): e00155-19. https://doi.org/10.1128/iai. 00155-19 8. Donegan NP, Manna AC, Tseng CW et al (2019) CspA regulation of Staphylococcus aureus carotenoid levels and σ(B) activity is controlled by YjbH and Spx. Mol Microbiol 112(2):532–551. https://doi.org/10.1111/ mmi.14273 9. Liu Y, Wu N, Dong J et al (2010) SsrA (tmRNA) acts as an antisense RNA to regulate Staphylococcus aureus pigment synthesis by base pairing with crtMN mRNA. FEBS Lett 584 (20):4325–4329. https://doi.org/10.1016/j. febslet.2010.09.024 10. Lan L, Cheng A, Dunman PM et al (2010) Golden pigment production and virulence gene expression are affected by metabolisms in Staphylococcus aureus. J Bacteriol 192 (12):3068–3077. https://doi.org/10.1128/ jb.00928-09 11. Singh VK (2014) Lack of a functional methionine sulfoxide reductase (MsrB) increases oxacillin and H2O2 stress resistance and enhances pigmentation in Staphylococcus aureus. Can J Microbiol 60(9):625–628. https://doi.org/ 10.1139/cjm-2014-0360 12. Sapp AM, Mogen AB, Almand EA et al (2014) Contribution of the nos-pdt operon to virulence phenotypes in methicillin-sensitive Staphylococcus aureus. PLoS One 9(9):e108868. https://doi.org/10.1371/journal.pone. 0108868 13. Hammer ND, Schurig-Briccio LA, Gerdes SY et al (2016) CtaM is required for menaquinol

oxidase aa3 function in Staphylococcus aureus. MBio 7(4):e00823-16. https://doi.org/10. 1128/mBio.00823-16 14. Clauditz A, Resch A, Wieland KP et al (2006) Staphyloxanthin plays a role in the fitness of Staphylococcus aureus and its ability to cope with oxidative stress. Infect Immun 74 (8):4950–4953. https://doi.org/10.1128/ iai.00204-06 15. Liu GY, Essex A, Buchanan JT et al (2005) Staphylococcus aureus golden pigment impairs neutrophil killing and promotes virulence through its antioxidant activity. J Exp Med 202(2):209–215. https://doi.org/10.1084/ jem.20050846 16. Young AJ, Lowe GL (2018) Carotenoidsantioxidant properties. Antioxidants 7(2). https://doi.org/10.3390/antiox7020028 17. Mishra NN, Liu GY, Yeaman MR et al (2011) Carotenoid-related alteration of cell membrane fluidity impacts Staphylococcus aureus susceptibility to host defense peptides. Antimicrob Agents Chemother 55(2):526–531. https:// doi.org/10.1128/aac.00680-10 18. Tiwari KB, Gatto C, Wilkinson BJ (2018) Interrelationships between fatty acid composition, staphyloxanthin content, fluidity, and carbon flow in the Staphylococcus aureus membrane. Molecules 23(5):1201. https:// doi.org/10.3390/molecules23051201 19. Chamberlain NR, Mehrtens BG, Xiong Z et al (1991) Correlation of carotenoid production, decreased membrane fluidity, and resistance to oleic acid killing in Staphylococcus aureus 18Z. Infect Immun 59(12):4332–4337. https:// doi.org/10.1128/iai.59.12.4332-4337.1991 20. Xiong Z, Kapral FA (1992) Carotenoid pigment levels in Staphylococcus aureus and sensitivity to oleic acid. J Med Microbiol 37 (3):192–194. https://doi.org/10.1099/ 00222615-37-3-192 21. Yang Y, Wang H, Zhou H et al (2020) Protective effect of the golden staphyloxanthin biosynthesis pathway on Staphylococcus aureus under cold atmospheric plasma treatment. Appl Environ Microbiol 86(3). https://doi. org/10.1128/aem.01998-19 22. Pannu MK, Hudman DA, Sargentini NJ et al (2019) Role of SigB and staphyloxanthin in radiation survival of Staphylococcus aureus. Curr Microbiol 76(1):70–77. https://doi. org/10.1007/s00284-018-1586-x 23. Chen F, Di H, Wang Y et al (2016) Smallmolecule targeting of a diapophytoene desaturase inhibits S. aureus virulence. Nat Chem Biol 12(3):174–179. https://doi.org/10.1038/ nchembio.2003

S. aureus Pigment Quantification 24. Gao P, Davies J, Kao RYT (2017) Dehydrosqualene desaturase as a novel target for antivirulence therapy against Staphylococcus aureus. MBio 8(5):e01224-17. https://doi.org/10. 1128/mBio.01224-17 25. Kahlon AK, Roy S, Sharma A (2010) Molecular docking studies to map the binding site of squalene synthase inhibitors on dehydrosqualene synthase of Staphylococcus aureus. J Biomol Struct Dyn 28(2):201–210. https://doi.org/ 10.1080/07391102.2010.10507353

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26. Liu CI, Liu GY, Song Y et al (2008) A cholesterol biosynthesis inhibitor blocks Staphylococcus aureus virulence. Science 319(5868):1391–1394. https://doi.org/10.1126/science.1153018 27. Morikawa K, Maruyama A, Inose Y et al (2001) Overexpression of sigma factor, sigma(B), urges Staphylococcus aureus to thicken the cell wall and to resist beta-lactams. Biochem Biophys Res Commun 288(2):385–389. https:// doi.org/10.1006/bbrc.2001.5774

Chapter 2 Analysis of Murein Hydrolases and Proteases Through Zymography Nichole A. Seawell and Jeffrey L. Bose Abstract Zymography has been used to analyze enzymatic activity and processing of enzymes for many years. We have used bacterial cells copolymerized into the acrylamide gel to analyze specific activity of murein hydrolases of interest. In addition, this method has been widely used to examine and distinguish protease activities using different substrates. This chapter provides instruction for zymography of both extracellular murein hydrolases and proteases produced by Staphylococcus aureus. Key words Murein hydrolase, Protease, Zymography, Proteolytic cleavage, In-gel digestion

1

Introduction The principle behind zymography is relatively straight forward: Protein separation by SDS-PAGE in a gel that is polymerized with the enzyme’s substrate, followed by renaturation of the enzyme in the gel, and visualization of its activity by degradation of the polymerized substrate [1, 2]. This technique has been used extensively for decades for assessing murein hydrolase activity. One benefit of this assay compared to other assays is that proteolytic processing of the murein hydrolases can be observed as long as the processed form remains catalytically active. Another benefit of this type of assay is the ease of screening various conditions. For example, replicate samples can be loaded in multiple lanes of the same zymography gel, separated by SDS-PAGE, each lane cut out, and placed in different renaturing conditions [1, 2]. This allows one to screen for inhibitors, temperature dependencies, and metal cofactors, just to name a few (compare panels A and C in Fig. 1). In addition, for dual-function enzymes, a sample can be loaded onto two different gel substrates and examined. A hallmark example of this is the use of Micrococcus lysodeikticus or Staphylococcus aureus cells to look at the activities of the glucosaminidase or amidase

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Murein hydrolase zymography of AtlA from S. aureus supernatants. Panels contain either Staphylococcus (a and c) or Micrococcus (b and d) cells to show activity of amidase and glucosaminidase, respectively. Panels (c) and (d) contain EDTA in the renaturation buffer, demonstrating that using a metal chelator inhibits the Zn-containing amidase, but not glucosaminidase

activity, respectively, of the Atl murein hydrolase (compare panels A and B in Fig. 1). This tool has been invaluable when confirming enzymatically inactive Atl variants [3]. In addition to murein hydrolases, zymography has been used as a simple and useful method to study other enzymes such as proteases. Much like murein hydrolases, the substrate used, and renaturation buffer conditions depends on the protease of interest. Commonly used substrates include gelatin and casein, and some companies make these in precast gels that are commercially available. Renaturation of the zymography gel in the presence of specific protease inhibitors has been useful in determining the likely identification of proteases in a complex mixture or when proteases are of similar size, as well to identity protease processing events [4, 5]. We have routinely examined both S. aureus supernatant proteins as well as cell-associated proteins using these methodologies.

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Below, we provide methods for collecting samples, producing the gels, renaturing the proteins, and how best to visualize the results.

2

Materials

2.1 Preparation of S. aureus Culture Supernatants

1. Filter-sterilized culture supernatant samples for testing (see Note 1). 2. 15-mL conical tubes. 3. Amicon Ultra-4 3000 MWCO filter unit. 4. Refrigerated benchtop centrifuge with 15-mL tube adapters.

2.2 Preparation of S. aureus Cell Wall Fractions

1. Cell pellet samples for testing (see Note 1). 2. 3 M LiCl. 3. 3-mL 10,000 MWCO Slide-A-Lyzer. 4. KPO4 buffer: 1 M K2HPO4 and 1 M KH2PO4 in water, adjusted to pH 7.2. 5. 15-mL conical tubes. 6. Refrigerated shaking incubator. 7. Refrigerated benchtop centrifuge with 15-mL tube adapters. 8. 10-mL syringe with 18-gauge needle. 9. Amicon Ultra-4 3000 MWCO filter unit.

2.3

Zymography

1. H2O. 2. 40% (w/v) acrylamide (see Note 2). 3. 1.5 M Tris–HCl pH 8.8. 4. 1.0 M Tris–HCl pH 6.8. 5. 10% (w/v) Sodium dodecyl sulfate (SDS). 6. 10% (w/v) Ammonium persulfate (APS): Make a fresh solution for each experiment or store at 4  C for 1 week. 7. TEMED. 8. Substrate: (a) For murein hydrolase: Micrococcus or Staphylococcus cells. (b) For protease: Casein or Gelatin. 9. Glass plates for acrylamide gels. 10. Acrylamide gel comb. 11. 100% ethanol (200 proof) or isopropanol. 12. Reagent(s) to determine protein concentration. For example, Bradford Reagent or Bicinchoninic Acid (BCA) Protein Assay. 13. Commercially available SDS Sample Buffer. 14. Heat block at 60  C.

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15. Pre-stained protein standard ladder. 16. 10 Electrophoresis Buffer: 30.3 g Tris Base, 188 g glycine, 10 g SDS, 800 mL water, adjust to pH 8.3 then add water to a final volume of 1 L. 17. Renaturation Buffer: (a) For murein hydrolases: 1% Triton X-100 in 25 mM Tris– HCl, pH 8.0. (b) For proteases: 2.5% Triton X-100. 18. Development buffer (for proteases): 50 mM Tris, 200 mM NaCl, 5 mM anhydrous CaCl2, and 0.02% Brij 35, adjusted to pH 7.5. 19. Platform shaker. 20. Incubator set at 37  C. 21. Stain solution: (a) For murein hydrolases: 1% (w/v) Methylene blue in 0.01% (w/v) KOH. (b) For proteases: 40% (v/v) Methanol, 10% (v/v) acetic acid and 0.5% (w/v) Coomassie blue R-250. 22. Destain solution: (a) For murein hydrolases: water. (b) For proteases: 40% (v/v) methanol and 10% (v/v) acetic acid. 23. Kim wipes. 24. White light source for imaging.

3

Methods

3.1 Preparation of S. aureus Culture Supernatants

1. Add 4 mL of filter-sterilized culture supernatant into an Amicon Ultra-4 3000 MWCO filter unit. 2. Centrifuge at 3160  g at 4  C for 30 min. 3. After centrifugation, 400–600 μL should be retained in the filter unit (~eightfold concentrated compared to original sample volume). If a more concentrated sample is required, remove the flow-through and repeat centrifugation (see Notes 3 and 4). 4. Quantify the protein concentration of each sample using Bradford or BCA assay. 5. Store samples at

3.2 Preparation of S. aureus Cell Wall-Associated Proteins (See Note 5)

80  C until ready to use.

1. Resuspend cell pellet in 5 mL of 3 M LiCl. 2. Incubate at 10  C with shaking at 250 rpm for 10 min in a conical tube laying on its side. 3. Centrifuge at 3160  g at 4  C for 10 min.

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4. Remove 4 mL of supernatant. 5. Using a syringe, add 3 mL of supernatant to a 3-mL 10,000 MWCO Slide-A-Lyzer. 6. Dialyze overnight in 0.01 M KPO4 buffer at room temperature. 7. Remove dialyzed sample and add to Amicon Ultra-4 3000 MWCO filter unit. 8. Centrifuge at 3160  g for 30 min at 4  C. 9. 400–600 μL should be retained in the filter unit. If a more concentrated sample is required, remove the flow-through and repeat centrifugation (see Notes 3 and 4). 10. Samples can be frozen at 3.3 Casting Zymogram Substrate Gel

80  C until ready to use.

1. Mix components for the resolving gel without adding TEMED, following the recipe in Table 1, which is based on a BioRad MiniPROTEAN 1 mm system. 2. Add substrate to resolving gel solution as follows (see Note 6): (a) Standard concentration of protease substrate in the resolving gel solution is 0.1% (w/v) for casein and 0.05% (w/v) for gelatin. These concentrations are similar to the precast BioRad zymography gels that were previously available for purchase (personal communication). (b) Standard concentration of murein hydrolase substrate in the resolving gel solution is ~2  109 CFU/mL for heatkilled S. aureus cells (see Note 7) and 1 mg/mL for commercially available Micrococcus lysodeikticus cells (see Note 8). 3. Add TEMED. 4. Pour gel in between glass plates, up to approximately 75% the height of the shorter plate. 5. Carefully pipette ethanol or isopropanol on top of the gel (see Note 9). 6. After gel has polymerized (approximately 30 min, see Note 10) remove the ethanol or isopropanol and rinse with water. 7. Mix components for the stacking gel, following the recipe in Table 1. 8. Pour stacking gel slowly onto the resolving gel. 9. Immediately insert a gel comb into the stacking gel. 10. Wait for stacking gel to polymerize (approximately 30 min).

3.4 SDS-PAGE and Zymography

1. Using water, normalize the samples to be loaded so that they are at the same concentration and volume. 2. Add the appropriate amount of SDS Sample Buffer to each sample tube.

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Table 1 Recipes for 5% stacking gel and 12% resolving gel components of a 1.0-mm zymogram gel cast Stacking gel

Resolving gel

H2O

2.2 mL

H2O

3.45 mL

40% acrylamide (ratio)

375 μL

40% acrylamide

2.4 mL

1.0 M Tris pH 6.8

380 μL

1.5 M Tris pH 8.8

2.0 mL

10% SDS

30 μL

10% SDS

80 μL

10% APS (fresh)

30 μL

10% APS (fresh)

80 μL

TEMED

3 μL

TEMED

3.2 μL

3. Heat samples at 60  C for 15 min. 4. Load samples into wells of gel along with a pre-stained protein ladder. 5. Run gel at 100 V for ~1.5 h (see Note 11). 6. Once gel is done running remove gel from plates and place in a square petri dish. 7. Rinse gel for 30 min on a platform shaker at room temperature with enough Renaturation Buffer to cover the gel. 8. Decant Renaturation Buffer. 9. Cover gel in more buffer and cover container (see Notes 12 and 13). (a) For murein hydrolases: Renaturation Buffer. (b) For proteases: Development solution. 10. Incubate statically at 37  C overnight. 11. After overnight incubation rinse with 10 mL of water. 12. Repeat this rinse two more times to remove any buffer. 13. Stain the gel as appropriate for each substrate (see Note 14). (a) For murein hydrolases: cover gel in water and add ~1 mL of staining solution. (b) For proteases: staining solution without dilution. 14. Shake on platform shaker at room temperature for 30 min or until gel is darkly stained. 15. Destain gel with several washes of destaining solution appropriate for each substrate as described in Subheading 2, at room temperature on a platform shaker (see Note 15). 16. Image using a white light source.

Staphylococcus aureus Zymography

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Notes 1. The protocol presented in this chapter is based on analysis of culture supernatant samples harvested from stationary-phase S. aureus cultures grown in tryptic soy broth at 37  C and 250 rpm. However, murein hydrolase activity could also be analyzed in the cell wall-associated fraction, and protease activities analyzed in cell membrane or cytosolic fractions. 2. The ratio of acrylamide to methylene:bisacrylamide can be altered to control for the pore size in order to obtain greater resolution of the protein of interest. We typically use 40% acrylamide/Bis 29:1 solution from BioRad to produce a 12% gel. 3. If a more concentrated protein is desired after 30 min, pipette the sample up and down a few times in the concentrator tube in order to resuspend the proteins that have moved toward the filter and centrifuge again in 10-min intervals. 4. Concentrating proteins might also concentrate proteases leading to undesired consequences by protein degradation. We have tested if adding proteases inhibitors before concentrating the samples had an effect on Atl murein hydrolase activity and saw no noticeable effect. 5. Cell wall-associated proteins can also be collected in alternative ways including boiling the cells in SDS, extraction using 6 M urea, or using membrane extraction methods, depending on specific need. 6. For murein hydrolase zymography, it is easiest to resuspend the cell substrate in the acrylamide gel mixture before adding the SDS to reduce the amount of bubbles when mixing. If performed carefully, this is not a big concern and the presence of SDS may improve suspension of the substrate. 7. To prepare heat-killed S. aureus cells as a substrate: (a) Dilute overnight culture to an OD600 of 0.1 into 100 mL of TSB in a 500-mL flask. (b) Grow at 37  C and 250 rpm for 3 h. (c) Serially dilute a sample of the culture to determine the CFU/mL of the culture. (d) Autoclave cells using a 30-min liquid cycle. (e) Cool to room temperature. (f) Place cells at 4  C until the next day when CFU counts can be used to calculate the CFU/mL. (g) Concentrate cells by centrifugation at 3160  g for 15 min. (h) Resuspend the cells to a total volume of 450 μL.

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(i) Cells can be stored at 4  C. Do not freeze or the cells will not suspend in the gel. 8. Micrococcus lysodeikticus and S. aureus cells are also commercially available from Millipore-Sigma. Any other bacteria cells could be used in this step as a substrate. We typically use S. aureus cells in the range of 109–1010 total cells per gel. Hybrid gels with a mixture of S. aureus and Micrococcus cells as substrate can also be used. 9. Adding 100% (v/v) ethanol or isopropanol right after pouring the resolving gel will remove any bubbles and creates an even surface onto which the stacking gel can be poured. 10. The time it takes for polymerizing the gel will vary depending on the nature and concentration of substrate within the gel but does take longer than a normal acrylamide gel without substrate added. 11. Running the gel takes longer than it normally would for a gel without substrate and the length of time can vary depending on the substrate used. 12. Wrapping the container in foil or plastic wrap will prevent the reaction buffer from evaporating overnight in the incubator. 13. Keep in mind that different proteases may behave differently at different pH and temperatures. 14. We have found staining is almost essential for protease zymography. For murein hydrolases, staining may improve visualizing bands; however, we have noticed that with a quality camera and appropriate background color this step may be unnecessary. 15. The gel will destain faster if you place a folded Kimwipe in the dish while shaking to soak up the dye. Periodically remove Kimwipe and destain solution and replace with a new until the bands in the gel are clear and the background is still dark. References 1. Fernandez-Resa P, Mira E, Quesada AR (1995) Enhanced detection of casein zymography of matrix metalloproteinases. Anal Biochem 224 (1):434–435 2. Leber TM, Balkwill FR (1997) Zymography: a single-step staining method for quantitation of proteolytic activity on substrate gels. Anal Biochem 249(1):24–28. https://doi.org/10. 1006/abio.1997.2170 3. Bose JL, Lehman MK, Fey PD et al (2012) Contribution of the Staphylococcus aureus Atl AM and GL murein hydrolase activities in cell division, autolysis, and biofilm formation. PLoS

One 7(7):e42244. https://doi.org/10.1371/ journal.pone.0042244 4. Rice K, Peralta R, Bast D et al (2001) Description of staphylococcus serine protease (ssp) operon in Staphylococcus aureus and nonpolar inactivation of sspA-encoded serine protease. Infect Immun 69(1):159–169. https://doi. org/10.1128/IAI.69.1.159-169.2001 5. Sieprawska-Lupa M, Mydel P, Krawczyk K et al (2004) Degradation of human antimicrobial peptide LL-37 by Staphylococcus aureus-derived proteinases. Antimicrob Agents Chemother 48 (12):4673–4679. https://doi.org/10.1128/ AAC.48.12.4673-4679.2004

Chapter 3 Detection and Quantification of Secreted Nuclease Activity in Staphylococcus aureus Culture Supernatants Richard E. Wiemels, Rebecca A. Keogh, and Ronan K. Carroll Abstract Staphylococcal secreted nuclease contributes to S. aureus virulence by degrading neutrophil extracellular traps (NETs), which allows the bacterium to evade the host immune system and has also been shown to promote biofilm dispersal. In this chapter, two methods for detecting nuclease activity are described, both of which have increased sensitivity compared to the traditional nuclease agar method. Key words Staphylococcus, Nuclease, Virulence factor, Salmon sperm DNA, FRET

1

Introduction Staphylococcal nuclease (Nuc, also called thermonuclease, micrococcal nuclease) is a well-studied secreted virulence factor of Staphylococcus aureus [1–3]. Initially, purification and characterization of Nuc [4] led to studies of the protein’s folding due to the presence of a proline–peptide bond that must be in the cis-conformation to allow the protein to fold correctly [5]. Nuc has also been studied in the context of protein secretion due to (a) the presence of an unusually long secretion signal sequence, and (b) because it is initially secreted as a longer, less active form (NucB) that is subsequently processed to a shorter, more active form (NucA) via an unknown mechanism [6]. Secretion systems of other species of bacteria have also been studied using Nuc as a reporter [7, 8]. The biological role of Nuc in S. aureus has been linked to immune evasion [1–3]. Specifically, Nuc degrades neutrophil extracellular traps (NETs), allowing bacteria to evade killing and exclude macrophages from abscesses [3]. Nuc also has a role in dispersing biofilms by facilitating detachment of cells and allowing for the bacteria to spread to additional sites [6, 9, 10]. Detecting secreted Nuc activity has traditionally involved preparing a type of nuclease agar and checking for zones of clearing

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Salmon sperm DNA degradation assay. The salmon sperm DNA degradation assay was performed using culture supernatants from three independent cultures of wild-type S. aureus and a ppiB mutant (a ppiB mutant strain is known to exhibit decreased nuclease activity [11]). Culture supernatants were incubated with salmon sperm DNA and subsequently separated by electrophoresis on 1% agarose. Sterile TSB broth was used as a negative control. Following electrophoresis, densitometry was performed on each lane using ImageJ. The resulting plots were graphed in Microsoft Excel and the area of peak intensity for each sample is indicated. The ppiB mutant samples peak is located higher up the gel compared to the wild type indicating that most of the DNA fragments are higher in molecular weight. This indicates less nuclease activity in the ppiB culture supernatants

around colonies, indicating digestion of DNA in the media. Here we describe two alternate methods of screening that can capture subtler differences in activity. The first method involves incubating culture supernatants with high molecular weight salmon sperm DNA, separating the samples at various time points on agarose gels, and observing differences in DNA degradation [11] (Fig. 1). The second method monitors the digestion of an oligonucleotide probe labeled on the 50 end with a fluorophore and the 30 end with a quencher [8]. When the oligonucleotide is intact, fluorescence of the probe is quenched (Fig. 2). In the presence of Nuc, the oligonucleotide is cleaved, releasing the fluorophore from the quencher and resulting in an increase in fluorescence (Fig. 2). Nuc activity is proportional to the amount of fluorescence observed, so the fluorescence detected can be used to compare the activity between strains [11] and quantify Nuc activity [8].

Nuclease Activity Assays

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19

Materials

2.1 Preparation of Cell-Free Supernatants

1. Supernatants from synchronized cultures (see Note 1). 2. 0.22 μm PVDF syringe-filters. 3. 10 ml syringes. 4. Benchtop centrifuge. 5. Tryptic soy broth (TSB): 3% (w/v) TSB in H2O. Sterilize by autoclaving.

2.2 Salmon Sperm DNA Degradation Assay

1. High molecular weight salmon sperm DNA solution: 10 mg/ ml dissolved in sterile H2O (see Note 2). 2. 0.2 ml nuclease-free PCR tubes. 3. Nuclease-free H2O. 4. 1% (w/v) agarose gel. 5. Ethidium bromide solution: 1 mg/ml ethidium bromide in H2O. 6. 6 DNA loading dye: 10 mM Tris–HCl (pH 7.6) 0.03% bromophenol blue, 0.03% xylene cyanol FF, 60% glycerol 60 mM EDTA. 7. 1 Tris-Acetate-EDTA (TAE): 40 mM Tris, 40 mM acetate, 1 mM EDTA pH 8.3. 8. Thermocycler. 9. Multichannel pipette capable of pipetting 25–50 μl volumes. 10. Horizontal electrophoresis apparatus and power supply. 11. Gel-imaging system containing a UV transilluminator.

2.3 Densitometry Analysis of Degraded Salmon Sperm DNA

1. Computer with ImageJ and graphing software.

2.4 FRET Probe Nuclease Assay

1. FRET substrate: An oligonucleotide (50 -CCC CGG ATC CAC CCC-30 ) modified on the 50 end with Cy3 and the 30 end with Blackhole quencher 2 (BHQ2) (see Note 3) [6, 12]. 2. FRET assay buffer: 20 mM Tris pH 8.0, 10 mM CaCl2. 3. Temperature controlled 96-well plate reader capable of excitation at 552 nm and emission at 580 nm (see Note 4). 4. White or black 96-well plates (see Note 5).

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Fig. 2 FRET probe nuclease assay. The FRET probe nuclease assay was performed using culture supernatants from three independent cultures of wild-type S. aureus and TSB broth as a negative control. Culture supernatants were diluted with sterile TSB and incubated with the FRET probe in a Synergy HTX 96-well plate reader at 37  C. An increase in fluorescence was observed from 0 to 50 min, after which no further increase in signal was detected. No increase in fluorescence was observed in the TSB negative control

3

Methods

3.1 Preparation of Cell-Free Supernatants

1. Centrifuge 10 ml of each growth-phase synchronized S. aureus culture to be analyzed at the desired time point at 3000  g for 15 min to pellet cells. 2. Syringe-filter each supernatant through 0.22 μm PVDF (low protein binding) filters. 3. Dilute supernatants in sterile TSB. For the salmon sperm DNA degradation assay, start with diluting supernatants 1:10 and adjust accordingly.

3.2 Salmon Sperm DNA Degradation Assay

1. Dissolve salmon sperm DNA in nuclease-free water to 10 mg/ml. 2. Aliquot 50 μl to 0.2 ml tubes. Set up one tube each for 5, 10, 15, 20, 25 and 30 min incubations. In parallel, for each timepoint setup a negative control tube containing DNA and sterile TSB alone (i.e., a no nuclease control). 3. Add 50 μl of diluted supernatant to the tubes containing salmon sperm DNA. If possible, perform this step for each

Nuclease Activity Assays

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reaction tube simultaneously using a multichannel pipette (see Note 6). Pipette up and down four to five times to mix and place tubes in a thermocycler set to 37  C. 4. At each time point, remove the corresponding tube, add 20 μl of 6X DNA loading dye to the mixture, and pipette up and down to stop the reaction. 5. Load 5 μl of each tube to wells of a 1% (w/v) agarose gel and run gel electrophoresis using 1 TAE buffer for 60 min at 100 V. 6. Stain the agarose gel containing electrophoresed DNA using ethidium bromide. Ethidium bromide can be added to the gel at a final concentration of 1 μg/ml before pouring the gel, or the gel can be post-stained in a solution of 1 μg/ml ethidium bromide. 7. Visualize and document gel image on a UV transilluminator and gel-imaging system (Fig. 1) (see Note 7). 3.3 Densitometry Analysis of Degraded Salmon Sperm DNA

1. To quantify the differences in nuclease activity, densitometry analysis can be performed using ImageJ software (or similar densitometry analysis software) on each sample in each lane (see Note 8). 2. Open the image of the gel in ImageJ and draw a box around the first lane (make sure the box goes all the way from the very top of the image to the very bottom). 3. Go to Analyze ! Plot profile. 4. Click the “Live” button on the plot window. 5. Click the “More” button on the plot window and select “Plot options.” 6. Select the “Fixed y-axis scale” and “Vertical profile” buttons. 7. Click the “Save” button on the plot window and save the resulting file. 8. Go back to the gel image and drag the selection rectangle to the second lane to be analyzed. 9. Click the “Save” button on the plot window and save the resulting file. 10. Repeat this process until the data has been saved from all lanes. 11. The resulting files contain the XY coordinates for each plot. 12. Open each file in Microsoft Excel (or another graphing program) and combine into one graph (as shown in Fig. 1) (see Note 9).

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3.4 FRET Probe Nuclease Assay

1. For the cell-free supernatant prepared in Subheading 3.1 which will serve as the comparison for all samples being assayed (i.e., the wild-type strain), perform a dilution series of cell-free supernatant in a 25 μl volume in row A of a 96-well plate (see Note 10). 2. Dilute the FRET substrate to 2 μM in assay buffer (20 mM Tris pH 8.0, 10 mM CaCl2) and add 30 μl to each well of row B that corresponds to dilution series in row A. 3. Preheat the plate in the microplate reader at 37  C for 5 min. 4. Using a multichannel pipette, pipette 25 μl of the probe (in row B) into the diluted supernatants (in row A). Pipette up and down twice to mix and immediately start reading fluorescence (excitation at 552 nm and emission at 580 nm). 5. As the probe is degraded, an increase in fluorescence is observed (see Note 11). 6. Identify an appropriate dilution of the supernatant where the reaction takes 30–60 min to run to completion. 7. Repeat this procedure for all samples using this same dilution factor (i.e., wild-type and mutant strains) (Fig. 2). 8. If desired, the amount of Nuc activity can be calculated using a standard curve generated from dilutions of purified Nuc [6].

4

Notes 1. In our lab, typical starter cultures of S. aureus are grown overnight at 37  C and 250 rpm in 3 ml TSB. The following day, these cultures are diluted 1:100 in 10 ml of fresh TSB and grown for 3 h to mid-exponential phase. The mid-exponential phase cultures are then diluted into 25 ml TSB (in a 250 ml conical flask) to an OD600 of 0.05. Supernatants are harvested after 15 h of growth. If an earlier time point is chosen, less Nuc will accumulate in the media and less dilution will be necessary to assay for activity. 2. High molecular weight DNA from a variety of sources should work in this assay. Avoid purchasing salmon sperm DNA that is sold as a blocking reagent for Southern and northern blots as it is typically already sheared to 200–500 bp in length. Salmon sperm nuclei work well as the genomic DNA is entirely intact. 3. Suppliers of oligonucleotides provide these modifications for Taqman probe applications. The probe can be suspended in water to 100 μM and stored at 20  C in 40 μl aliquots. 4. The assay can also be performed at room temperature if the plate reader does not have a heating function. However, it is better to heat the reaction to a set temperature to ensure reproducible results as ambient temperatures can vary.

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5. Clear plates can also be used, but the fluorescence generated in one well can influence the results in adjacent wells. 6. A multichannel pipette is very useful for timing this assay and is essential for the FRET probe assay. 7. The qualitative results of this assay can be observed in the gel image. As the salmon sperm DNA is cleaved by nuclease, the DNA has the appearance of a “smear” in the lane rather than a discrete band. The intensity of the smear varies throughout the lane and a region of peak intensity is typically observed (see Fig. 1). A difference in apparent molecular weight of the region of peak intensity indicates differences in nuclease activity. No DNA degradation should be observed in the TSB control, and therefore one discrete band corresponding to undigested DNA should remain. 8. Some gel imagers are equipped with image analysis software that can generate a histogram of the intensity of fluorescence over a cross-section of gel area. Alternatively, densitometry analysis can be performed with freely available software such as ImageJ. 9. The region of peak intensity for each sample is apparent. Differences in the peaks between two samples indicate a difference in nuclease activity (compare WT to ppiB in Fig. 1). 10. Due to the sensitivity of the FRET probe nuclease assay, it is necessary to start with a dilution series to find a suitable working range. 11. The reaction proceeds very quickly. At higher concentrations of supernatant, the reaction will run to completion within seconds. It is important to identify a suitable dilution of supernatant whereby the reaction can be monitored over 30–60 min.

Acknowledgments This work was supported in part by grant AI128376 from the National Institute of Allergy and Infectious Diseases. References 1. Olson ME, Nygaard TK, Ackermann L et al (2013) Staphylococcus aureus nuclease is an SaeRS-dependent virulence factor. Infect Immun 81(4):1316–1324. https://doi.org/ 10.1128/iai.01242-12 2. Schilcher K, Andreoni F, Uchiyama S et al (2014) Increased neutrophil extracellular trap-mediated Staphylococcus aureus clearance through inhibition of nuclease activity by

clindamycin and immunoglobulin. J Infect Dis 210(3):473–482. https://doi.org/10. 1093/infdis/jiu091 3. Thammavongsa V, Missiakas DM, Schneewind O (2013) Staphylococcus aureus degrades neutrophil extracellular traps to promote immune cell death. Science 342(6160):863–866. https://doi.org/10.1126/science.1242255

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4. Tucker PW, Hazen EE Jr, Cotton FA (1978) Staphylococcal nuclease reviewed: a prototypic study in contemporary enzymology. I. Isolation; physical and enzymatic properties. Mol Cell Biochem 22 (2–3):67–77. https://doi.org/10.1007/ bf00496235 5. Tucker PW, Hazen EE Jr, Cotton FA (1979) Staphylococcal nuclease reviewed: a prototypic study in contemporary enzymology. IV. The nuclease as a model for protein folding. Mol Cell Biochem 23(3):131–141. https://doi. org/10.1007/bf00219452 6. Kiedrowski MR, Kavanaugh JS, Malone CL et al (2011) Nuclease modulates biofilm formation in community-associated methicillinresistant Staphylococcus aureus. PLoS One 6 (11):e26714. https://doi.org/10.1371/jour nal.pone.0026714 7. Poquet I, Ehrlich SD, Gruss A (1998) An export-specific reporter designed for grampositive bacteria: application to Lactococcus lactis. J Bacteriol 180(7):1904–1912. https:// doi.org/10.1128/jb.180.7.1904-1912.1998 8. Recchi C, Rauzier J, Gicquel B et al (2002) Signal-sequence-independent secretion of the

staphylococcal nuclease in Mycobacterium smegmatis. Microbiology 148(Pt 2):529–536. https://doi.org/10.1099/00221287-148-2529 9. Mann EE, Rice KC, Boles BR et al (2009) Modulation of eDNA release and degradation affects Staphylococcus aureus biofilm maturation. PLoS One 4(6):e5822. https://doi.org/ 10.1371/journal.pone.0005822 10. Moormeier DE, Bose JL, Horswill AR et al (2014) Temporal and stochastic control of Staphylococcus aureus biofilm development. MBio 5(5):e01341-01314. https://doi.org/ 10.1128/mBio.01341-14 11. Wiemels RE, Cech SM, Meyer NM et al (2017) An intracellular peptidyl-prolyl cis/trans isomerase is required for folding and activity of the Staphylococcus aureus secreted virulence factor nuclease. J Bacteriol 199(1):e00453-16. https://doi.org/10.1128/jb.00453-16 12. Lee SP, Porter D, Chirikjian JG et al (1994) A fluorometric assay for DNA cleavage reactions characterized with BamHI restriction endonuclease. Anal Biochem 220(2):377–383. https://doi.org/10.1006/abio.1994.1353

Chapter 4 Quantitative Hemolysis Assays Miranda J. Ridder, Seth M. Daly, Pamela R. Hall, and Jeffrey L. Bose Abstract Many strains of Staphylococcus aureus produce a variety of cytolysins that target many different cell types to both fight the immune system and acquire nutrients. This includes hemolysins which destroy erythrocytes and are well studied virulence factors. Traditionally, hemolysin activity is measured on blood agar plates due to the simplicity of the assay. While this is telling, it cannot encapsulate the full story because S. aureus is known to behave differently in broth and on agar. Furthermore, plate-based assays are primarily semiquantitative and often a more accurate determination of hemolytic potential is needed to discern differences between strains. Here, we describe a method to quantify hemolysin activity from broth or similarly grown cells. Key words Hemolysin, Toxin, Alpha-hemolysin, Staphylococcus aureus, Red blood cell lysis

1

Introduction Staphylococcus aureus produces and secretes several different types of hemolysins during infection. While they can kill many types of cells, hemolysins are noted for their ability to lyse red blood cells. Beta-hemolysin is encoded by hlb and functions as an Mg2+-dependent sphingomyelinase [1]. The effects of β-hemolysin are best observed on sheep’s blood at colder temperatures after warm incubation, and complete hemolysis does not occur until blood-agar plates are placed in 4  C after incubation at 37  C [1]. Of note, some strains are β-hemolysin deficient due to a phage insertion. S. aureus also produces δ-hemolysin, which is encoded by hld within RNAIII. Gamma-hemolysin is encoded by hlgABC, forms two bi-component pore-forming proteins, HlgAB and HlgCB, and has not yet been shown to have species specificity [2]. Perhaps one of the best characterized S. aureus hemolysins is α-hemolysin, a heptameric pore-forming β-barrel toxin encoded by hla. Secreted α-hemolysin can lyse red blood cells, some leukocytes, and epithelial cells by binding to the host receptor ADAM10, a zincdependent metalloprotease [3]. Rabbit blood is particularly

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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susceptible to α-hemolysin because rabbit red blood cells express higher levels of ADAM10 when compared to humans and other mammals [4]. It should be noted that the nomenclature for these proteins is diverse. For example, α-hemolysin is also often called Hla as well as α-toxin. In addition to these hemolysins, S. aureus produce cytotoxic phenol soluble modulins (PSMs). S. aureus possesses four co-transcribed α-PSMs and two β-type PSMs [5, 6]. These small peptides have a variety of effects on both bacterial and eukaryotic cells, and are known to lyse human red blood cells [7]. Notably, δ-toxin is considered a PSM due to its size and structure similarity to α-PSMs [5, 6]. Hemolysis assays have traditionally made use of nutrient agar plates incorporating blood in the medium. These semiquantitative assays are simple to perform and are a great tool to analyze the presence or absence of hemolysin production. While this is truly telling of hemolytic activity, it is difficult to quantify all but large differences as well as the kinetics of hemolysin production. Another disadvantage of blood-agar plates is that they represent a very different growth condition than in liquid culture and gene expression may not be similar between the two environments. To determine hemolysin activity more precisely, and from non-surface cultured cells, quantitative hemolysis assays provide a powerful tool for analyzing activity of these important secreted toxins. This protocol is based on an existing protocol published in 1988 [8].

2

Materials 1. Growth medium of choice. 2. Blood (see Notes 1 and 2). 3. Clear flat-bottom 96-well plates. 4. 15-ml conical tubes. 5. 1 phosphate buffered saline (1 PBS): 0.8% (w/v) sodium chloride, 0.02% (w/v) potassium chloride, 0.14% (w/v) disodium phosphate, 0.02% (w/v) potassium dehydrate phosphate pH 7.4. Store at 4  C. 6. Microplate reader for optical density measurements at 650 nm. 7. 1 PBS containing 1% (v/v) Triton X-100. 8. Single channel pipettes and tips capable of dispensing 100–300μl volumes. 9. Multichannel pipette and tips capable of dispensing 200μl volumes. 10. Refrigerated benchtop centrifuge.

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11. 37  C static incubator. 12. S. aureus samples to be tested for hemolysin activity (see Note 3).

3

Methods

3.1 Preparation of Blood Solution

1. Mix bottle of blood by gentle inversion. 2. Add 12 ml of cold 1 PBS to a 15-ml conical tube. 3. Slowly add 1 ml of blood to the 1 PBS and mix by gently inverting (see Note 4). 4. Centrifuge at 1450  g for 10 min at 4  C. 5. Remove supernatant and discard (see Note 5). 6. Wash again with 12 ml of cold 1 PBS with gentle mixing. 7. Centrifuge at 1450  g for 10 min at 4  C. 8. Resuspend in 12 ml of cold PBS, yielding a 4% (v/v) concentration.

3.2 Prepare Microtiter Plate with Test Samples (Plate 1 in Fig. 1)

1. Add 150μl of PBS to all sample wells except the first and last columns. 2. Add 300μl of undiluted sample to the first column in duplicate or triplicate (see Fig. 1—Plate 1). 3. Perform twofold serial sample dilution by transferring 150μl of undiluted supernatant from column 1 to column 2. 4. Repeat the transfer of 150μl from column 2 to column 3 and continue across the plate, leaving the last column blank.

3.3 Prepare Microtiter Plate with Blood (Plate 2 in Fig. 1)

1. Add 100μl of prepared blood from step 8 of Subheading 3.1 to wells of a new 96-well plate, except for two wells in the last column, which will serve as blank well controls (see Fig. 1— Plate 2). 2. Transfer 100μl of each test sample (from Plate 1) onto Plate 2. Mix gently (see Note 6). 3. In the last column: (a) Add 100μl of 1 PBS to 3 wells containing blood. These will serve as blood only (negative) control wells. (b) Add 100μl of 1 PBS with 1% Triton X-100 to 3 wells containing blood. These will serve as your complete lysis (positive) control wells. (c) Add 200μl of 1 PBS with Triton X-100 to the 2 wells without blood to serve as blank wells. 4. Cover plate with lid or adhesive seal (see Note 7). 5. Place in 37  C incubator (static) for 1 h.

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Fig. 1 Plate 1 is the layout for dilution of hemolysin samples with duplicate samples depicted. Diluted samples are transferred to Plate 2 containing red blood cells. Plate 2 also shows control samples in column 12

6. After 1 h, use multichannel pipette or plate shaker to gently mix and resuspend any red blood cells that may have settled to the bottom of the plate.

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7. Read at OD650 nm on a microplate reader (see Note 8). The OD650 nm measures the density of erythrocytes in each well, so the higher the value, the less hemolysis (more cells) present. A low OD indicates more hemolysis (less cells). 8. To determine HA50, which is the dilution that results in 50% hemolysis: (a) Prepare an XY graph where X-values are the log of the dilution factor and Y-values are the OD values. (b) Fit the curves with a 4-parameter logistic regression. (c) The EC50 (the concentration that gives half-maximal lysis) calculated is the HA50. The HA50 between strains may then be compared.

4

Notes 1. The source of blood chosen should reflect the hemolysin of interest. For example, it is well known that rabbit red blood cells are most susceptible to α-hemolysin lysis, whereas human blood is more appropriate for detecting PSM hemolysis, and sheep blood works well for β-hemolysin. 2. Blood should be defibrinated or include an anti-coagulant. Typically, we purchase 50% (v/v) rabbit blood in Alsever’s solution (prevents coagulation) from Colorado Serum Company. The protocol assumes a 50% (v/v) stock concentration of blood to reach a final 4% (v/v) concentration of washed RBCs. 3. Often, these samples are supernatants from liquid mediagrown cells, but any source that allows the collection of proteins can be used. For example, we typically test either 0.2μm filtered bacterial supernatants or purified toxin. For comparison between bacterial strains we grow cultures and match their final OD600 nm to 0.05 with fresh media to account for growth differences. Cultures are then spun and supernatants passed through 0.2μm polyethersulfone filters. Any time point will work and should be chosen based on the timing of the hemolysin to be tested. 4. All mixing with blood should be performed gently whether by inversion or pipette since erythrocytes are easily lysed. 5. When removing supernatant from blood, use a pipette and do NOT pour off. Pouring will disrupt the blood cell pellet and decrease the final blood cell concentration. 6. By mixing 100μl of sample and 100μl of prepared blood in column 1, the lowest dilution obtained is 1:2. 7. Seals are recommended to prevent evaporation if greater than 1 h incubation times are needed.

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8. An alternative is to measure the heme released during red blood cell lysis. In this case, the plate should be centrifuged after the 1 h incubation, supernatants transferred to a new plate, and supernatant measurements performed at OD450 nm. This optical density measures the heme released and a high number correlates with increased cell lysis. References 1. Huseby M, Shi K, Brown CK et al (2007) Structure and biological activities of beta toxin from Staphylococcus aureus. J Bacteriol 189 (23):8719–8726. https://doi.org/10.1128/jb. 00741-07 2. Spaan AN, Vrieling M, Wallet P et al (2014) The staphylococcal toxins γ-hemolysin AB and CB differentially target phagocytes by employing specific chemokine receptors. Nat Commun 5:5438–5438. https://doi.org/10.1038/ ncomms6438 3. Berube BJ, Bubeck Wardenburg J (2013) Staphylococcus aureus alpha-toxin: nearly a century of intrigue. Toxins (Basel) 5(6):1140–1166 4. Wilke GA, Bubeck Wardenburg J (2010) Role of a disintegrin and metalloprotease 10 in Staphylococcus aureus alpha-hemolysin-mediated cellular injury. Proc Natl Acad Sci U S A 107 (30):13473–13478. https://doi.org/10.1073/ pnas.1001815107

5. Cheung GY, Joo HS, Chatterjee SS et al (2014) Phenol-soluble modulins—critical determinants of staphylococcal virulence. FEMS Microbiol Rev 38(4):698–719. https://doi.org/10. 1111/1574-6976.12057 6. Wang R, Braughton KR, Kretschmer D et al (2007) Identification of novel cytolytic peptides as key virulence determinants for communityassociated MRSA. Nat Med 13 (12):1510–1514. https://doi.org/10.1038/ nm1656 7. Cheung GY, Duong AC, Otto M (2012) Direct and synergistic hemolysis caused by Staphylococcus phenol-soluble modulins: implications for diagnosis and pathogenesis. Microbes Infect 14 (4):380–386. https://doi.org/10.1016/j. micinf.2011.11.013 8. Bernheimer AW (1988) Assay of hemolytic toxins. Methods Enzymol 165:213–217

Chapter 5 In Vitro Assay for Quantifying Clumping of Staphylococcus aureus Heidi A. Crosby, Jakub M. Kwiecinski, and Alexander R. Horswill Abstract Staphylococcus aureus interacts with fibrinogen in plasma to form macroscopic clumps of cells. A simple and rapid slide agglutination test using rabbit plasma has been employed in clinical labs to distinguish S. aureus from most coagulase-negative Staphylococci. The method described here is a quantitative clumping assay in which S. aureus cells are mixed with either plasma or purified fibrinogen, and clumps are allowed to sediment out of solution. Clearing of the overlying solution is monitored over time by measuring the optical density at 600 nm and comparing these values to the initial turbidity. This simple assay can be used to study regulation and expression of various cell wall-anchored adhesins. Key words Staphylococcus aureus, Clumping, Agglutination, Fibrinogen, Clumping factor

1

Introduction One of the hallmarks of S. aureus is its ability to bind to the plasma protein fibrinogen and form macroscopic aggregates, commonly referred to as clumping [1]. S. aureus clumping is mediated by two cell wall-anchored surface proteins termed clumping factors, ClfA and ClfB, that specifically bind to distinct regions within fibrinogen [2, 3] (reviewed in [4]). Fibrinogen is a symmetrical, propellershaped protein, and S. aureus interacts with the distal ends of the molecule, allowing fibrinogen to act as a bridge between neighboring cells [2]. The ability of S. aureus to bind fibrinogen is important in a number of infection settings, including endocarditis [5], sepsis [6], kidney abscesses [7], skin infections [8], and septic arthritis [9]. The slide agglutination test for clumping factor (also sometimes referred to as bound coagulase) has been used for several decades as a rapid diagnostic test for S. aureus. In the simplest version, a S. aureus colony is resuspended in sterile saline and mixed with a drop of rabbit or human plasma on a glass slide, and agglutination is scored within seconds. Newer versions of the slide

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Clumping of Staphylococci in the presence of human plasma. (a) S. aureus strain LAC log-phase cells were washed and resuspended in 1.5 ml of either buffer (PBS) or 1.25% human plasma. Tubes were allowed to stand at room temperature for 3 h. (b) Quantification of clumping in a variety of S. aureus and S. epidermidis strains after 2 h of incubation in the presence of 1.25% human plasma. Values represent averages and standard deviations of three independent experiments. S. epidermidis serves as a representative non-clumping coagulase-negative Staphylococcal species

test use latex beads coated with plasma or a combination of fibrinogen and IgG, which will detect both clumping factor and protein A [10, 11]. These tests are highly accurate (>95% accuracy with a range of S. aureus isolates [12]), although certain members of the coagulase-negative Staphylococci (CoNS), including S. lugdunensis and S. schleiferi, make clumping factors and can give positive results [13]. While these slide agglutination tests are sufficient for clinical applications, it is useful to have a quantitative measure of S. aureus clumping when comparing strains in a research setting. We have employed a simple clumping test in which S. aureus cells are incubated with either human plasma or fibrinogen, and clumps are allowed to settle to the bottom of the tube (Fig. 1). A reduction in turbidity of the overlying solution is monitored over time, and is used as a proxy to determine the relative amount of clumping, which can be readily compared between strains [14, 15]. This is a relatively simple assay that can be easily adapted to test for clumping in the presence of fibrinogen or plasma from a variety of sources.

2

Materials 1. Lyophilized human plasma (see Note 1). 2. 1.5 mg/ml human fibrinogen in 1
Phosphate-buffered saline (PBS) (see Note 2). 3. Brain Heart Infusion (BHI) medium.

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4. BHI agar plates. 5. Sterile 15 ml culture tubes. 6. Shaking incubator set at 37  C. 7. Plastic cuvettes (1.5 ml volume, 1.0 cm path length). 8. Spectrophotometer capable of reading optical density at 600 nm (OD600). 9. Sterile microcentrifuge tubes (2.0 ml size). 10. Microcentrifuge. 11. Sterile 1
PBS. 12. 96-well plate. 13. 96-well plate reader capable of reading OD600.

3

Methods 1. Streak bacterial strains from freezer stocks onto brain heart infusion (BHI) agar (see Note 3). Invert plates and grow overnight at 37  C. 2. For each bacterial strain, pick a single colony to inoculate 5 ml of brain heart infusion (BHI) medium in a culture tube. Grow overnight (~16 h) at 37  C with shaking (200–250 rpm). 3. The following morning, dilute the overnight cultures 1:100 each into 5 ml of fresh BHI medium. Grow at 37  C with shaking to an optical density (OD600) of 1.5 (see Note 4). 4. When the OD600 has reached 1.5 (see Note 5), transfer 1.5 ml of each culture to a 2 ml microcentrifuge tube. Centrifuge for 1 min at maximum speed (approximately 20,000
g) in a microcentrifuge and discard the supernatants. Resuspend each cell pellet in 1.5 ml of 1
PBS. 5. Centrifuge for 1 min at maximum speed and discard the supernatants. Resuspend each cell pellet in 1.5 ml of 1
PBS. 6. Add 18.75 μl of 100% (w/v) plasma or 1.5 mg/ml fibrinogen prepared as described in Notes 1 and 2 to each cell sample tube. Vortex each cell suspension briefly (~1 s) to mix and note the starting time (see Note 6). 7. Remove 100 μl from each cell suspension and use to measure the OD600 in a 96-well plate reader to get the OD600t ¼ 0. Discard the cells after taking the measurement. 8. Let the tubes prepared in step 6 (above) sit statically at room temperature and avoid disturbing them. At desired time points (see Note 7), carefully remove 100 μl from the top of each cell suspension and transfer to a 96-well plate to measure the

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Fig. 2 Clumping time courses with a range of fibrinogen (a) and plasma (b) concentrations. S. aureus strain LAC cells were incubated with the indicated concentrations of fibrinogen or plasma in PBS and clumping was quantified over 2 h

OD600 (see Note 8). This is recorded as the OD600t. An alternative measurement option is also presented (see Note 9). 9. Calculate the relative clumping for each time point (t) using the equation below (see Note 10): %clumping ¼ 100
½ðOD600t¼0  OD600t Þ=OD600t¼0 Þ: Representative data obtained with this assay is presented in Figs. 1 and 2.

4

Notes 1. Human plasma can be obtained from blood donors or purchased in lyophilized form from a variety of vendors. We have successfully used plasma from healthy donors that was mixed 1:1 with heparin/dextran sulfate to prevent coagulation. We have also used commercially available lyophilized human plasma that was pretreated with 4% trisodium citrate. If lyophilized plasma is used, reconstitute in distilled deionized water to make a 100% (w/v) solution, and store at 20  C in 1 ml aliquots for later use. 2. Fibrinogen can be purchased from various commercial sources. Prepare a 1.5 mg/ml solution in phosphate-buffered saline and freeze 1 ml aliquots at 20  C. Note that fibrinogen should be dissolved using gentle agitation rather than vortexing. 3. Other rich growth medium plates, such as tryptic soy agar (TSA) or sheep blood agar plates, can also be used at this step. 4. To measure the optical density, remove a small amount of the culture with a sterile pipette (e.g., 300 μl) and dilute with two volumes of sterile BHI. The mixture is transferred to a disposable plastic cuvette (1.0 cm path length) and the optical density is measured at 600 nm using a spectrophotometer, after

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blanking the instrument with sterile BHI. The reading is multiplied by the dilution factor (in this case three) to get the OD600 of the culture. 5. Some strains of S. aureus, particularly mutants, may grow more slowly than others. To compare clumping of different strains it is best to grow them so that they have reached the same optical density. We recommend resuspending cell pellets from each culture in PBS (as described in the step 4 of the protocol), and then storing the resuspended cells at room temperature until all strains are at the same time point or phase of growth before proceeding to step 5. 6. We typically use plasma at a final concentration of 1.25% (w/v) and fibrinogen at a concentration of ~18.5 μg/ml, although a range of concentrations will produce similar results after 2 h of incubation (Fig. 2). However, we find that high concentrations of plasma (>5%) can interfere with the assay, likely because the high viscosity impedes settling of clumps. Fibrinogen concentrations in plasma range from 1.5 to 4 mg/ml, and thus a 1.25% plasma solution would contain 18.75–50 μg/ml of fibrinogen. 7. It is recommended that time point measurements be taken every 30 min for 2 h, although the spacing and number of time points can be adjusted depending on the behavior of the strains being tested. 8. For consistency, the same time points should be measured each time an experiment is repeated, because removing even a small volume from the tube can affect later measurements. 9. It is also possible to take continuous readings without sacrificing assay volume using a turbidity meter that will hold small test tubes and can measure the absorbance near the top of the column of liquid. 10. We typically repeat the experiment three times on different days, calculate the % aggregated for the desired time points, and then average those values and calculate standard deviations. For most S. aureus strains, the bulk of the clumping occurs in the first 1–2 h and comparing the 2-h time points is often optimal for observing variations between different strains.

Acknowledgments H. Crosby was supported by American Heart Association postdoctoral fellowship 15POST25720016. J. M. Kwiecinski was supported by a Swedish Society for Medical Research postdoctoral fellowship and by American Heart Association postdoctoral fellowship 17POST33670580. Research in the lab of A. R. Horswill was

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supported by a Merit Award (I01 BX002711) from the Department of Veteran Affairs, project 3 of NIH grant AI083211, and NIH public health service grants AI141490 and AI153185. References 1. Much H (1908) A preliminary stage of the fibrin enzymes in cultures of Staphylococcus aureus. Biochem Z 14:143–155 2. Hawiger J, Timmons S, Strong DD et al (1982) Identification of a region of human fibrinogen interacting with staphylococcal clumping factor. Biochemistry 21 (6):1407–1413 3. Walsh EJ, Miajlovic H, Gorkun OV et al (2008) Identification of the Staphylococcus aureus MSCRAMM clumping factor B (ClfB) binding site in the alphaC-domain of human fibrinogen. Microbiology 154(Pt 2):550–558. https://doi.org/10.1099/mic.0.2007/ 010868-0 4. Crosby HA, Kwiecinski J, Horswill AR (2016) Staphylococcus aureus aggregation and coagulation mechanisms, and their function in hostpathogen interactions. Adv Appl Microbiol 96:1–41. https://doi.org/10.1016/bs.aambs. 2016.07.018 5. Moreillon P, Entenza JM, Francioli P et al (1995) Role of Staphylococcus aureus coagulase and clumping factor in pathogenesis of experimental endocarditis. Infect Immun 63 (12):4738–4743 6. Josefsson E, Higgins J, Foster TJ et al (2008) Fibrinogen binding sites P336 and Y338 of clumping factor A are crucial for Staphylococcus aureus virulence. PLoS One 3(5):e2206. https://doi.org/10.1371/journal.pone. 0002206 7. Cheng AG, Kim HK, Burts ML et al (2009) Genetic requirements for Staphylococcus aureus abscess formation and persistence in host tissues. FASEB J 23(10):3393–3404. https:// doi.org/10.1096/fj.09-135467 8. Kwiecinski J, Jin T, Josefsson E (2014) Surface proteins of Staphylococcus aureus play an important role in experimental skin infection. APMIS

122(12):1240–1250. https://doi.org/10. 1111/apm.12295 9. Josefsson E, Hartford O, O’Brien L et al (2001) Protection against experimental Staphylococcus aureus arthritis by vaccination with clumping factor A, a novel virulence determinant. J Infect Dis 184(12):1572–1580. https://doi.org/10.1086/324430 10. Essers L, Radebold K (1980) Rapid and reliable identification of Staphylococcus aureus by a latex agglutination test. J Clin Microbiol 12 (5):641–643 11. Dickson JI, Marples RR (1986) Coagulase production by strains of Staphylococcus aureus of differing resistance characters: a comparison of two traditional methods with a latex agglutination system detecting both clumping factor and protein A. J Clin Pathol 39(4):371–375 12. Lairscey R, Buck GE (1987) Performance of four slide agglutination methods for identification of Staphylococcus aureus when testing methicillin-resistant staphylococci. J Clin Microbiol 25(1):181–182 13. Freney J, Brun Y, Bes M et al (1988) Staphylococcus lugdunensis sp. nov. and Staphylococcus schleiferi sp. nov., two species from human clinical specimens. Int J Syst Bacteriol 38 (2):168–172 14. Walker JN, Crosby HA, Spaulding AR et al (2013) The Staphylococcus aureus ArlRS two-component system is a novel regulator of agglutination and pathogenesis. PLoS Pathog 9(12):e1003819. https://doi.org/10.1371/ journal.ppat.1003819 15. Crosby HA, Schlievert PM, Merriman JA et al (2016) The Staphylococcus aureus global regulator MgrA modulates clumping and virulence by controlling surface protein expression. PLoS Pathog 12(5):e1005604. https://doi.org/10. 1371/journal.ppat.1005604

Chapter 6 Measuring Staphylococcal Promoter Activities Using a Codon-Optimized β-Galactosidase Reporter Christina N. Krute, Nichole A. Seawell, and Jeffrey L. Bose Abstract The lacZ gene and corresponding β-galactosidase enzyme has been a mainstay for bacterial reporter systems for decades. We have used this versatile reporter to analyze expression profiles from strains grown both on solid media and from broth culture. The standard broth protocol can also be adapted for a 96-well plate to allow high-throughput screening of promoter reporter constructs under a variety of conditions. Furthermore, codon-optimization of the E. coli lacZ gene has greatly improved activity levels of β-galactosidase in S. aureus, facilitating improved sensitivity for screening assays, detection of low-activity promoters, and use of small sample volumes. In this chapter, details are provided for both standard and high-throughput quantitative assays that we have routinely used for S. aureus transcriptional profiling. Key words lacZ, Staphylococcus, Reporter, Miller units, Promoter fusion

1

Introduction Regulation of bacterial gene expression has been examined for decades using a variety of techniques and technologies. One of the most convenient approaches to study gene expression is through the use of easily detected reporter constructs. These reporters can be constructed in a variety of ways including simple transcriptional promoter reporter fusions used to examine expression kinetics, coupled with promoter truncations or modifications to interrogate promoter regulation. In addition, when using translational reporter fusions with native ribosome binding sites, one can infer both transcriptional and translational differences (see Note 1). At the heart of all these designs is the need for both an easy-todetect and quantifiable output. Over the years, reporter genes have been chosen based on detection needs, experimental considerations, and accessibility. These reporters often take the shape of directly detectable outputs including fluorescence (i.e., genes encoding fluorescent proteins), and luminescence (i.e., firefly or bacterial luciferase genes), or indirect measurements of reporter

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gene enzymatic activity (i.e., genes encoding β-galactosidase or chloramphenicol acetyltransferase). One mainstay of bacterial reporter genes used for many bacteria is the lacZ gene, encoding β-galactosidase. The use of lacZ was pioneered by Dr. Jeffrey Miller when he worked out the kinetics of β-galactosidase using 2-Nitrophenyl β-D-galactopyranoside (ONPG), giving us the guidelines of proper activity range and calculation, with the results reported as Miller units [1] (see Note 2). Miller units ¼ 1000 

ðOD420 nm  ð1:75  OD550 nm ÞÞ time ð min Þ  volume ðmlÞ  OD600 nm

The use of lacZ and β-galactosidase lies partially in substrate availability and utility. Enzymatically, β-galactosidase can recognize several important substrates leading to different outputs. Some of the more commonly used substrates are summarized in Table 1. In addition to those listed in Table 1, additional substrates are available for β-galactosidase with detection using fluorescence, luminescence, or antibiotic resistance (see Note 3). One complicating factor involved in reporter design is creating constructs to measure promoter activities that are lowly expressed or are poorly translated due to non-canonical ribosome binding sites. Previously, in another book in this series, we demonstrated the value of optimized ribosome binding sites [2]. This is a valuable way to amplify a weak signal as long as native translation is not necessary. In addition, we recently produced codon-optimized genes for fluorescent reporters in Staphylococcus species [3]. The lacZ gene of E. coli has a high (56%) GC content compared to S. aureus (33%). To aid in our studies, we had the E. coli lacZ codon-optimized for S. aureus by decreasing the GC content to 36% and removing rare codons [4]. When comparing expression from the same promoter in S. aureus, this codon-optimized lacZ yields 16-fold higher activity compared to non-codon optimized E. coli lacZ (see Note 4 and Fig. 1). Below, we provide two methodologies to determine β-galactosidase activity: (1) a high-throughput method for rapid screening and (2) a more traditional method for increased accuracy of activity determination. Of note, we have found the highthroughput method to be less accurate, but useful for screening a large set of conditions. Our lab then follow-ups those results with the traditional method.

2 2.1

Materials General Supplies

1. S. aureus strain harboring lacZ reporter fusion construct. 2. Clear flat-bottom 96-well plates.

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Table 1 Commonly used β-galactosidase substrates Substrate

Detection

Primary use

4-methyumbelliferyl-β-D-galactopyranoside (MUG)

Fluorescence (365 nm λex; 445 nm λem)

Agar or liquid

2-Nitrophenyl β-D-galactopyranoside (ONPG)

Colorimetric (yellow)

Liquid assay

5-bromo-4-chloro-3-indolyl-D-galactopyranoside (X-gal)

Colorimetric (blue)

Agar

Fig. 1 S. aureus RN4220 with transcriptional hla promoter fusion (Phla) driving expression of either the original E. coli lacZ (lacZ) or the codon-optimized lacZ (colacZ). Data are representative of four independent experiments each with three biological replicates; error bars are standard error of the mean

3. Spectrophotometer (420–600 nm range). 4. Spectrophotometer cuvettes. 5. Bead beater homogenizer. 6. 1.7 ml microcentrifuge tubes. 7. Microcentrifuge. 8. 2.0 ml screw cap tubes. 9. Glass beads (0.1 mm diameter). 10. Z-Buffer: 60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM β-mercaptoethanol, pH 7.0 (see Note 5). 11. 2-Nitrophenyl β-D-galactopyranoside (ONPG) stock solution: 4 mg/ml dissolved in water. 12. Stop solution: 1 M sodium carbonate (Na2CO3). 13. Reagent(s) to determine protein concentration. 2.2 Supplies Specific to High-Throughput Method

1. Microplate reader with absorbance range of 420–600 nm. 2. Multichannel (20–200μl range) pipette.

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3. Triton Z-Buffer: 60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM β-mercaptoethanol, pH 7.0, 0.5% (v/v) Triton X-100 (see Note 5). 4. Benchtop centrifuge with rotor for 96-well plates.

3

Methods

3.1 High-Throughput Screening Method

1. Grow reporter fusion strain to desired growth phase. 2. Standardize reporter fusion strain to an OD600 nm ¼ 0.05 in a well of a clear flat-bottom 96-well plate to final volume of 200μl (see Note 6). 3. Incubate the 96-well plate shaking at 37  C for desired length of time or as appropriate for the experimental design. 4. Read final OD600 nm using a plate reader. 5. Pellet the cells by centrifugation of the 96-well plate at 200  g for 5 min and remove supernatant. 6. Resuspend the cells in 200μl of Triton Z-Buffer. 7. Disrupt the cells by pipetting up and down several (~10) times. 8. Remove 20μl to a new well containing 125μl Z-Buffer (see Note 7). 9. Add 29μl of 4 mg/ml ONPG. 10. Incubate the 96-well plate statically at 37  C, inspecting it periodically for formation of yellow color. 11. Once the sample appears slightly yellow (OD420 nm < 1.0), record the duration of incubation, and stop the reaction by adding 42μl Stop solution (see Note 8). 12. Read OD420 nm using a plate reader. 13. Calculate Miller units using the following formula: Miller units ¼ 1000 

OD420 nm time ð min Þ  volume ðmlÞ  OD600 nm

In the formula above, OD420 nm ¼ absorbance of yellow o-nitrophenol formation, time ¼ reaction time (minutes), volume ¼ volume (ml) of assayed sample, and OD600 nm ¼ optical density of well. 3.2 Modified Traditional Method

1. Grow reporter fusion strain to desired growth phase. 2. Pellet 1 ml of the strain by centrifugation at 21,000  g for 5 min (see Note 9). 3. Resuspend the cell pellet in 1.2 ml of Z-Buffer. 4. Transfer the resuspension to a 2.0 ml screw cap tube containing glass beads.

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5. Transfer the tube to a bead beater homogenizer and use manufacturer’s recommended setting for disruption of S. aureus cells (see Note 10). 6. Pellet the cell debris by centrifugation at 21,000  g for 5 min. 7. Transfer 100μl of each supernatant generated in step 6 to a new 1.7 ml tube. Set aside for protein quantification (step 13 below). 8. Transfer 5–100μl of the supernatant to a new 1.7 ml tube. Final volume should be adjusted to 700μl with addition of Z-Buffer (see Note 7). 9. Add 140μl of 4 mg/ml ONPG. 10. Incubate the sample statically at 37  C. 11. Once the sample appears slightly yellow (OD420 nm < 1.0), record the duration of incubation, and stop the reaction by adding 200μl 1 M Na2CO3. 12. Transfer the sample to a spectrophotometer cuvette and read at OD420 nm. 13. Calculate the protein concentration of your sample (see Note 11). 14. Calculate Modified Miller units (specific activity) as the following: Modified Miller units ¼ 1000 

OD420 nm time ð min Þ  volume ðmlÞ  protein concentration ðmg=mlÞ

In the formula above, OD420 nm ¼ absorbance of yellow onitrophenol formation, time ¼ reaction time (minutes), volume ¼ volume (ml) of assayed sample, and protein concentration ¼ concentration (mg/ml) of assayed sample.

4

Notes 1. It is important to note the slight differences between transcriptional and translational reporters. Transcriptional reporters primarily report promoter activity; thus, these reporters need to include native sigma factor binding sites, transcriptional start sites and any known binding sites for regulatory proteins. This does allow flexibility in other factors such as a non-native Shine Dalgarno site or modified first codon. For example, if a gene starts with GTG, then this site could be modified to ATG. These adjustments will amplify output of the reporter from the promoter due to better translation. In the case of a translational reporter, all of the same elements for transcriptional reporters are needed, but the native Shine Delgarno, complete

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50 UTR, and start codon should be used to best represent expression. Ideally, the plasmid used to produce the reporter should maximize flexibility. The plasmid that we developed, pJB185, has convenient cloning sites that allow for the use or removal of an optimized ribosome binding site with enhancer region to minimize secondary structure (Fig. 2). Furthermore, an XbaI site in-frame with lacZ makes generating translational reporters simple. Importantly, XbaI-cleaved DNA can be ligated to XbaI-, NheI-, or AvrII-digested DNA products. 2. In the Miller equation, the OD420 nm reading measures the yellow color from o-nitrophenol released from ONPG cleavage, 1.75  OD550 nm corrects for scatter from cell debris, time is the length of the assay in minutes, volume is the amount of sample used in the assay, and OD600 nm is the original optical density of the culture and is used to correct activity for bacterial numbers. In our experience, with proper centrifugation, the 1.75  OD550 nm correction is not needed. 3. We suggest the website www.markergene.com for substrate options. 4. For the data in Fig. 1, the hla promoter was amplified from pJB1017 [4] using primers JB22 (CCGCTAGCCATTTTCAT CATCCTTCTATTTTTTAAAACG) [4] and JBKU94 ( CAG GATCCTTGTACTGTTTGATATGGAACTCCTG), digested with BamHI and XbaI, and ligated into the same sites of pJB185 (colacZ, [4]) or pJB1020 (E. coli BL21 lacZ, unpublished) to create pNS07 and pNS08, respectively. 5. The rest of the Z-buffer can be made and stored at room temperature, but add fresh β-mercaptoethanol, pH 7.0, before use. 6. Account for addition of any other substances (e.g., antibiotics, chemical stressors, etc.) for final volume of 200μl. 7. The volume of cell lysate required for detection of β-galactosidase activity needs to be determined empirically for the individual strain, promoter, and/or growth/stress condition under study. The end goal is to have a final OD420 nm 0.25–1.0 (Miller stated ideally 0.6–0.9) as measured in a spectrophotometer with a 1 cm path length. However, this range could be longer depending on the spectrophotometer used. For example, the xenon lamp-based spectrophotometer in our lab allows for accuracy of readings well above OD420 nm ¼ 1.0. Readings for a plate reader should be individualized for the machine and sensitivity of detection, but is minimal concern for a high-throughput screen. This is also a function of time, so adjustments can be made to volume used or length of assay. We try to keep our assay times between 5 and 30 min.

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Fig. 2 Schematic of pJB185, a self-replicating plasmid designed for generating transcriptional or translational reporters. Sequence shown starts with position 1 of the plasmid and is the multi-cloning site (MCS) with restriction endonuclease sites marked. The highlighted ATG identifies the start codon of lacZ while the double underline shows the TIR optimized ribosome binding site. Other unique restriction endonuclease sites are shown on the map. CmR indicates location of gene encoding chloramphenicol resistanance in Staphylococcus species while AmpR show location of gene encoding ampicillin resistance for E. coli

8. The addition of stop solution will immediately increase the “yellowness” of the sample. 9. Cell pellets can be frozen at 80  C until use. 10. Settings on a MP Fastprep 24-5G for disruption of S. aureus cells is 2 cycles of 6.0 m/s speed for 40 s separated by a 300 s pause time. 11. A Bradford Protein Assay may be used to calculate protein concentration of the sample from a standard curve generated from a protein of known concentration. We prefer to use protein as our standardization rather than OD600 nm since it is more accurate and will compensate for any inefficient cell lysis during the bead beating step.

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Acknowledgment This work was supported in part by NIH grant AI121073 to JLB. References 1. Miller JH (1972) Experiments in molecular genetics. Cold Spring Harbor Laboratory, New York 2. Bose JL (2014) Genetic manipulation of staphylococci. In: Fey PD (ed) Staphylococcus epidermidis: methods and protocols, vol 1106. Methods in molecular biology. Humana Press, Totowa, pp 101–111. https://doi.org/10.1007/978-162703-736-5_8

3. Bose JL, Fey PD, Bayles KW (2013) Genetic tools to enhance the study of gene function and regulation in Staphylococcus aureus. Appl Environ Microbiol 79(7):2218–2224. https:// doi.org/10.1128/AEM.00136-13 4. Krute CN, Rice KC, Bose JL (2017) VfrB is a key activator of the Staphylococcus aureus SaeRS two-component system. J Bacteriol 199(5): e00828-16. https://doi.org/10.1128/JB. 00828-16

Chapter 7 Evaluation of Staphylococcus aureus Antibiotic Tolerance Using Kill Curve Assays Jessica N. Brandwein and Kelly C. Rice Abstract This chapter describes the use of antibiotic kill curves to examine the tolerance of Staphylococcus aureus to any antibiotic of interest. This is done by treating cultures with a super-minimum inhibitory concentration (MIC) of antibiotic and measuring viability over time by colony-forming units (CFUs). Kill curves provide a unique insight into S. aureus antibiotic tolerance and death patterns that may not be clear from other experiments, such as traditional MIC or Kirby–Bauer assays. Key words Antibiotic tolerance, Cell death, Persistence, MIC, CFU

1

Introduction In 2017, methicillin-resistant Staphylococcus aureus (MRSA) accounted for 323,700 estimated infections in hospitalized patients, 10,600 deaths, and $1.7 billion in estimated attributable healthcare costs [1]. Gaining heritable genetic antibiotic resistance such as gene(s) encoding efflux pumps, enzymes that inactivate the antibiotic or modify its cellular target, allows S. aureus to continue to grow at antibiotic concentrations that would otherwise inhibit or kill a cell lacking the resistance mechanism [2]. Genetic antibiotic resistance is also characterized by an increase in the minimum inhibitory concentration (MIC; lowest concentration of antibiotic that will inhibit planktonic growth in defined period of time) [3, 4]. Antibiotic treatment of infections caused by S. aureus and other notable pathogens can also be complicated by the phenomenon of antibiotic tolerance, whereby physiological and/or genetic changes decrease a bacterial cell’s susceptibility to the killing action of an antibiotic without a change in the MIC [5]. In other words, tolerant bacteria can survive at what would otherwise be lethal antibiotic concentrations (even at concentrations well above the MIC), but do not actively grow [2, 3, 5]. Specific examples of

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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genetic and physiological mechanisms governing bacterial antibiotic tolerance have been well discussed in several excellent review articles [2, 3, 5–7]. Phenotypic tolerance is often linked to the bacterial growth state, illustrated by the well-known phenomenon that slow-or non-growing cells are more tolerant to killing by β-lactam antibiotics relative to fast-growing cells [8–10]. In the case of S. aureus, its ability to grow as a biofilm and as a small colony variant (SCV) have both been linked to increased antibiotic tolerance (reviewed in [6, 11]). Cells in biofilms are often less susceptible to antibiotics not only due to the physical barriers inherent to the biofilm structure, but also due to slower growth and/or altered metabolism of cells occupying the deeper biofilm layers where oxygen is limited [2]. In both planktonic and biofilm cultures, even if the entire cell population is not tolerant to an antibiotic, a subpopulation of antibiotic tolerant cells known as “persisters” can form, which are thought to contribute to the difficulty of resolving chronic biofilm infections that are recalcitrant to antibiotic treatment [12]. These cells represent a highly antibiotic tolerant subpopulation of cells that emerge stochastically and are otherwise genotypically identical to the rest of the population [2, 7, 13]. This phenotype is also reversible and non-inheritable: Persister cells that are isolated from antibiotictreated cultures and then regrown in fresh medium lacking antibiotic, exhibit an identical pattern and magnitude of killing when re-exposed to that same antibiotic a second time [14, 15]. Antibiotic tolerance and persister cell formation continue to be active areas of research given their clinical importance, as much is still unknown about the underlying mechanistic basis for these phenomena. In S. aureus, the MazEF toxin-antitoxin system [16], cellular ATP depletion [17, 18], cyclic-di-AMP phosphodiesterase GdpP [19, 20], and the Cid/Lrg system [21, 22] have been associated with antibiotic tolerance and/or persister cell formation. Antibiotic resistance is commonly assessed by MIC using a planktonic-based assay, or by the agar plate-based Kirby–Bauer (disk diffusion) method [23]. The planktonic-based MIC assay involves exposing a defined number of bacterial cells in nutrient broth to antibiotic at different concentrations (typically in a twofold dilution series) and qualitatively assessing growth by the presence of turbidity [24, 25]. In this assay, MIC is defined as the lowest concentration of antibiotic that inhibits visible bacterial growth [24, 25]. For the Kirby–Bauer method, a bacterial suspension is spread evenly across a nutrient agar plate, and paper disks impregnated with antibiotics at defined concentrations are placed onto the surface. A “zone of inhibition,” observable as a clear ring of no bacterial growth around the disk, is evident after incubation if the bacteria are susceptible to the antibiotic [23].

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Although both the MIC and Kirby–Bauer methods provide clear insight into whether a bacterial strain is sensitive or resistant to a specific antibiotic, these assays are not able to capture information related to antibiotic tolerance. In contrast, the antibiotic kill curve assay described in this chapter is based on previous S. aureus publications [21, 22, 26] and examines bacterial viability over time when exposed to high (super-MIC) antibiotic concentrations. The observed patterns of bacterial cell killing can provide unique insights into both population-level antibiotic tolerance as well as formation of persister cell subpopulations. In brief, this antibiotic kill curve assay exposes exponentially growing S. aureus planktonic cultures to a super-MIC concentration of antibiotic, and the culture viability is monitored by serial dilution and colony-forming unit (CFU) plating over an 8-h period. As illustrated in Fig. 1, viability data can be presented as either CFU/ml (Fig. 1a) or expressed as a percentage of the CFU/ml when antibiotic is added to the culture (Fig. 1b), as this accounts for small differences in initial cell population size between replicates. This analysis can be used to compare antibiotic tolerance patterns between different S. aureus strains (such as wildtype and isogenic mutants as demonstrated in Fig. 1), or how different environmental growth conditions or compounds affect antibiotic tolerance.

2

Materials 1. Tryptic Soy agar (TSA) plates (round plates and square grid plates): 3% (w/v) Tryptic Soy medium, 1.5% (w/v) agar (see Note 1). 2. Tryptic soy broth (TSB): 3% (w/v) TSB. 3. TSB without glucose (TSB-G): 3% (w/v) tryptic soy broth (TSB) without glucose (see Note 2). 4. 80 mg/ml Vancomycin stock solution: Dissolve 800 mg of vancomycin in 10 ml of H2O, then filter-sterilize through a 0.2 μm syringe filter. Aliquot vancomycin solution and store at
20  C (see Note 3). 5. Sterile 10 ml luer lock syringe. 6. 0.2 μm syringe filter. 7. Sterile, round-bottomed culture tubes (15 ml volume). 8. Sterile 500 ml Erlenmeyer flasks with foiled tops. 9. Class 2AII biosafety cabinet. 10. Static plate incubator set at 37  C. 11. Shaking incubator set at 37  C, 250 rpm. 12. Spectrophotometer cuvettes (1.5 ml volume, 1.0 cm path length).

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Fig. 1 Antibiotic kill curve comparing tolerance of wild-type UAMS-1 and isogenic nos mutant strain (KR1010) to 80 μg/ml vancomycin. Vancomycin was added at “T ¼ 0”, cultures were grown for 8 h, and samples were taken at T ¼ 1, 2, 4, 6, and 8 for CFU and OD analysis. Data represent n ¼ 2 independent experiments, error bars ¼ standard deviation. (a) Data is represented as CFU/ml over time. (b) Data is represented as percent viability relative to the CFU/ml at time of vancomycin addition to culture

13. Spectrophotometer capable of reading optical density at 600 nm (OD600). 14. Sterile serological pipettes (5 ml, 50 ml volumes). 15. Single-channel pipettes (2–20 μl and 20–200 μl volumes). 16. Sterile tips for single-channel pipettes (2–20 μl and 20–200 μl volumes). 17. Benchtop vortexer. 18. Sterile microcentrifuge tubes (1.7 ml or 2 ml volumes).

S. aureus Kill Curve Assays

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Methods All steps below should be performed using standard aseptic technique and Biosafety Level 2 (BSL-2) precautions, including working under a Class 2AII biosafety cabinet as appropriate. 1. Streak S. aureus strain(s) to be analyzed onto TSA plates to obtain isolated colonies. Grow for 24 h in a static incubator at 37  C. 2. Inoculate a single isolated S. aureus colony from each plate in step 1 into a 15 ml culture tube containing 3 ml TSB. Incubate cultures overnight (14–16 h) at 37  C and 250 rpm (see Note 4). 3. Aliquot 40 ml of TSB-G each into sterile 500 ml Erlenmeyer flasks (1 flask per S. aureus overnight culture prepared in step 2 above) (see Note 5). Incubate flasks overnight at 37  C to check for sterility. 4. Vortex each overnight culture to disperse cells and measure the OD600 using cuvettes and a spectrophotometer. Calculate the volume of each culture needed to inoculate 40 ml of TSB-G to an initial OD600 ¼ 0.025. 5. Immediately remove a 1.5 ml sample (time of inoculation sample) from each culture in step 4. Set aside until needed in step 7 below. This timepoint is referred to as “T
2” on the graphs presented in Fig. 1. 6. Incubate all 40 ml cultures at 37  C and 250 rpm for 2 h (see Note 6). 7. While 40 ml cultures are incubating, measure the “time of inoculation” sample (set aside in step 5) OD600 using cuvettes and a spectrophotometer, and CFU/ml by setting up a 1:10 serial dilution series and CFU plating. In brief, add 50 μl of a culture sample into a microcentrifuge tube containing 450 μl sterile TSB. Vortex this dilution for 5 s, then immediately transfer 50 μl of this 10
1 dilution into a second tube containing 450 μl of sterile TSB. Repeat this procedure four more times for each culture flask, for a total of five dilution tubes (10
1 through 10
5) (see Note 7). Plate samples of the 10
3 through 10
5 dilution tubes in triplicate onto square TSA plates using the track plating method [27]. In brief, drop 10 μl aliquots each onto their own single grid at the top of a square TSA plate (two dilution tubes can be plated in triplicate onto one square TSA plate, see Fig. 2). Hold the plate at a 70 angle so that the droplets track down the surface of the agar toward the other edge. Once the tracks enter the grid at the bottom of the plate, place it flat on the bench, and allow the tracks to completely dry (15–20 min) (see Note 8). Invert each square plate (agar side on top) and incubate for 24 h overnight in a static incubator at 37  C.

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Fig. 2 Representative image of a square agar plate containing a bacterial culture diluted and plated as described in step 7 of Subheading 3

8. After 2 h of growth, remove culture flasks from the incubator. Remove 1.5 ml for CFU dilution plating and OD600 measurements as described in step 7 above. This timepoint, just before antibiotics are added, is referred to as “T ¼ 0” on the graphs presented in Fig. 1. 9. Add 40 μl of 80 mg/ml vancomycin stock solution to each culture, for a final concentration of 80 μg/ml (see Notes 9 and 10). 10. Incubate cultures for 8 more hours at 37  C, 250 rpm, removing 1.5 ml for CFU and OD600 analysis at 1, 2, 4, 6, and 8 h growth post-vancomycin addition. At each time point, immediately return the cultures to the shaking incubator after 1.5 ml volume is removed, and then proceed with OD600 and CFU plating as described in step 7 above. Incubate all CFU plates at 37  C for 24 h. 11. Count colonies from the dilution that yields 10–100 colonies per 10 μl track to determine CFU/ml at each time point. The CFU/ml for each sample can be calculated using the following formula (see Note 11): CFU=ml ¼ ðCFU  fold‐dilutionÞ=ðvolume platedÞ 12. To visualize kill curve data, the CFU/ml at each time point can be expressed as a percentage of the initial CFU/ml measured at T ¼ 0 (see step 8) and graphed with percent viability on the Yaxis and time on the X-axis (Fig. 1b). This standardizes for any slight differences in initial population size between samples before antibiotic is added to each culture.

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Notes 1. Tryptic Soy base contains the following components: Pancreatic digest of Casein 17 g/l, Papaic digest of Soybean 3 g/l, Dextrose 2.5 g/l, Sodium Chloride 5 g/l, Dipotassium Phosphate 2.5 g/l. Round agar plates should be used for streaking out frozen stock cultures, and square agar plates should be used for serial dilution and CFU plating during the assay itself. 2. This protocol is based on methodology used in [26], in which TSB without glucose (dextrose) was used to grow cultures in conditions promoting aerobic respiration (250 rpm, 1:12.5 volume to flask ratio). The exact growth medium, temperature, and culture aeration conditions should be modified to best reflect the physiological conditions relevant to your experiment. 3. For your antibiotic of interest, prepare stock solution at a concentration that is 1000-fold higher than the intended final concentration when added to each S. aureus culture. This will allow adding a small volume that will have minimal impact on the total culture volume. 4. The exact overnight culture incubation time can be adjusted for your S. aureus strain(s) of interest but should fall in the range of 12–18 h. It is important to keep this incubation time consistent for each experiment to minimize variability. 5. This 1:12.5 culture volume to flask ratio (40 ml of TSB in a 500 ml Erlenmeyer flask) is appropriate for promoting aerobic growth of S. aureus at 250 rpm. The volume of culture and size of Erlenmeyer flask used can be scaled up or down as needed if the culture volume to flask ratio is maintained. If aeration is a variable of interest to your experiment, this volume to flask ratio can be adjusted accordingly, keeping in mind that slower growing (less aerated) cultures in general will be more tolerant to certain antibiotics. 6. For cell-wall active antibiotics such as vancomycin, this experiment works best when antibiotic is added to early-mid exponential phase cultures. For S. aureus strain UAMS-1 (Fig. 1), 2 h aerobic growth in TSB-G represents this phase of growth. However, this time point may need to be optimized depending on the S. aureus strain used, growth conditions, and antibiotic of interest. If cultures are still in lag phase when antibiotic is added, cultures will be inherently more tolerant to antibiotic killing. 7. Adjust the dilution series for each bacterial strain and time point as needed, to achieve a 10–100 countable colony range from a 10 μl aliquot. It is best practice to plate three dilution

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tubes from each sample dilution series, one above and below the dilution tube expected to give this countable range. 8. It is important to let the 10 μl tracks dry completely before inverting the plate. If tracks are still wet, they may run together, and create a smear of bacterial growth instead of isolated countable colonies. For this same reason, it is very important to use square TSA plates that are completely dry on the agar surface. Excessive condensation/moisture on TSA plates can be removed by keeping plates at room temperature for several hours prior to use, or by drying plates under a sterile laminar flow (Class 2AII biosafety) cabinet. 9. Under the experimental conditions described in this protocol and Fig. 1, approximately 53 the MIC of vancomycin for S. aureus UAMS-1 is used, calculated using the previously published MIC for this strain [28, 29]. However, this protocol can be optimized for any specific antibiotic and/or S. aureus strain(s) of interest. It is recommended that kill curves be initially conducted with an antibiotic concentration at 10–30 the MIC of the wild-type S. aureus strain of interest, for which the MIC has been published or experimentally determined. The kill curve antibiotic concentration can be raised or lowered as required for increasing the sensitivity and dynamic range of killing observed over time. For example, if cultures experience rapid die-off (greater than 3-logs of killing relative to T ¼ 0) within the first 2 h, consider repeating the experiment with a reduced antibiotic concentration. 10. If comparing different wild-type strains (or wildtype and isogenic mutants) for the first time, the MICs should also be determined in a separate experiment to be sure that any differences in antibiotic tolerance observed in the kill curve assay are not due to inherent differences in the MIC. 11. For the track dilution method [27], the recommended countable colony range is 10–100. For calculating the CFU/ml, first multiply the number of counted colonies in a track by its corresponding fold-dilution. Then divide this number by the volume plated (for the track dilution method, this is 0.01 ml). For example, if 50 colonies are counted on a 10
3 dilution track, multiply 50 by 1000 (fold-dilution), then divide this number by 0.01 ml (volume plated), for a calculated 5  106 CFU/ml. The calculated CFU/ml for each technical replicate (triplicate replicates) should then be averaged.

Acknowledgment This work was supported in part by NIH grant AI118999 (KCR).

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References 1. CDC (2019) Antibiotic resistance threats in the United States, 2019. https://doi.org/10. 15620/cdc:82532 2. Brauner A, Fridman O, Gefen O et al (2016) Distinguishing between resistance, tolerance and persistence to antibiotic treatment. Nat Rev Microbiol 14(5):320–330. https://doi. org/10.1038/nrmicro.2016.34 3. Kester JC, Fortune SM (2014) Persisters and beyond: mechanisms of phenotypic drug resistance and drug tolerance in bacteria. Crit Rev Biochem Mol Biol 49(2):91–101. https://doi. org/10.3109/10409238.2013.869543 4. Mah TF (2014) Establishing the minimal bactericidal concentration of an antimicrobial agent for planktonic cells (MBC-P) and biofilm cells (MBC-B). J Vis Exp 83:e50854. https:// doi.org/10.3791/50854 5. Tuomanen E, Durack DT, Tomasz A (1986) Antibiotic tolerance among clinical isolates of bacteria. Antimicrob Agents Chemother 30(4):521–527. https://doi.org/10.1128/aac. 30.4.521 6. Bui LM, Conlon BP, Kidd SP (2017) Antibiotic tolerance and the alternative lifestyles of Staphylococcus aureus. Essays Biochem 61(1):71–79. https://doi.org/10.1042/EBC20160061 7. Yan J, Bassler BL (2019) Surviving as a community: antibiotic tolerance and persistence in bacterial biofilms. Cell Host Microbe 26(1):15–21. https://doi.org/10.1016/j.chom.2019.06.002 8. Tuomanen E, Cozens R, Tosch W et al (1986) The rate of killing of Escherichia coli by betalactam antibiotics is strictly proportional to the rate of bacterial growth. J Gen Microbiol 132(5):1297–1304. https://doi.org/10.1099/ 00221287-132-5-1297 9. Tuomanen E, Tomasz A (1990) Mechanism of phenotypic tolerance of nongrowing pneumococci to beta-lactam antibiotics. Scand J Infect Dis Suppl 74:102–112 10. Spoering AL, Lewis K (2001) Biofilms and planktonic cells of Pseudomonas aeruginosa have similar resistance to killing by antimicrobials. J Bacteriol 183(23):6746–6751. https:// doi.org/10.1128/JB.183.23.6746-6751.2001 11. Stokes JM, Lopatkin AJ, Lobritz MA et al (2019) Bacterial metabolism and antibiotic efficacy. Cell Metab 30(2):251–259. https:// doi.org/10.1016/j.cmet.2019.06.009 12. Lewis K (2005) Persister cells and the riddle of biofilm survival. Biochemistry (Mosc) 70(2):267–274. https://doi.org/10.1007/ s10541-005-0111-6

13. Lewis K (2010) Persister cells. Annu Rev Microbiol 64:357–372. https://doi.org/10. 1146/annurev.micro.112408.134306 14. Bigger JW (1944) Treatment of staphylococcal infections with penicillin—by intermittent sterilisation. Lancet 2:497–500 15. Keren I, Kaldalu N, Spoering A et al (2004) Persister cells and tolerance to antimicrobials. FEMS Microbiol Lett 230(1):13–18. https:// doi.org/10.1016/S0378-1097(03)00856-5 16. Ma D, Mandell JB, Donegan NP et al (2019) The toxin-antitoxin MazEF drives Staphylococcus aureus biofilm formation, antibiotic tolerance, and chronic infection. mBio 10(6): e01658-19. https://doi.org/10.1128/mBio. 01658-19 17. Conlon BP, Rowe SE, Gandt AB et al (2016) Persister formation in Staphylococcus aureus is associated with ATP depletion. Nat Microbiol 1:16051. https://doi.org/10.1038/nmicrobiol. 2016.51 18. Zalis EA, Nuxoll AS, Manuse S et al (2019) Stochastic variation in expression of the tricarboxylic acid cycle produces persister cells. MBio 10(5):e01930-19. https://doi.org/10. 1128/mBio.01930-19 19. Chung M, Borges V, Gomes JP et al (2018) Phenotypic signatures and genetic determinants of oxacillin tolerance in a laboratory mutant of Staphylococcus aureus. PLoS One 13(7):e0199707. https://doi.org/10.1371/ journal.pone.0199707 20. Griffiths JM, O’Neill AJ (2012) Loss of function of the gdpP protein leads to joint betalactam/glycopeptide tolerance in Staphylococcus aureus. Antimicrob Agents Chemother 56(1):579–581. https://doi.org/10.1128/ AAC.05148-11 21. Groicher KH, Firek BA, Fujimoto DF et al (2000) The Staphylococcus aureus lrgAB operon modulates murein hydrolase activity and penicillin tolerance. J Bacteriol 182(7):1794–1801. https://doi.org/10.1128/jb.182.7.1794-1801. 2000 22. Rice KC, Firek BA, Nelson JB et al (2003) The Staphylococcus aureus cidAB operon: evaluation of its role in regulation of murein hydrolase activity and penicillin tolerance. J Bacteriol 185(8):2635–2643. https://doi.org/10.1128/ jb.185.8.2635-2643.2003 23. Bauer AW, Perry DM, Kirby WM (1959) Single-disk antibiotic-sensitivity testing of staphylococci; an analysis of technique and results. AMA Arch Intern Med 104(2):208–216.

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https://doi.org/10.1001/archinte.1959. 00270080034004 24. Institute CaLS (2020) Performance standards for antimicrobial susceptibility testing, 30th edition vol CLSI document M100Ed30E. Wayne, PA 25. Wiegand I, Hilpert K, Hancock RE (2008) Agar and broth dilution methods to determine the minimal inhibitory concentration (MIC) of antimicrobial substances. Nat Protoc 3(2):163–175. https://doi.org/10.1038/nprot.2007.521 26. James KL, Mogen AB, Brandwein JN et al (2019) Interplay of nitric oxide synthase (NOS) and SrrAB in modulation of Staphylococcus aureus metabolism and virulence. Infect Immun 87(2):e00570-18. https://doi.org/ 10.1128/IAI.00570-18

27. Jett BD, Hatter KL, Huycke MM et al (1997) Simplified agar plate method for quantifying viable bacteria. BioTechniques 23(4):648–650. https://doi.org/10.2144/97234bm22 28. Weiss EC, Spencer HJ, Daily SJ et al (2009) Impact of sarA on antibiotic susceptibility of Staphylococcus aureus in a catheter-associated in vitro model of biofilm formation. Antimicrob Agents Chemother 53(6):2475–2482. https://doi.org/10.1128/AAC.01432-08 29. Meeker DG, Beenken KE, Mills WB et al (2016) Evaluation of antibiotics active against methicillin-resistant Staphylococcus aureus based on activity in an established biofilm. Antimicrob Agents Chemother 60(10):5688–5694. https://doi.org/10.1128/AAC.01251-16

Chapter 8 Fluorescence Polarization (FP) Assay for Measuring Staphylococcus aureus Membrane Fluidity Kiran B. Tiwari, Suranjana Sen, Craig Gatto, and Brian J. Wilkinson Abstract Fluorescence polarization is a method to determine membrane fluidity using a hydrophobic fluorescent dye that intercalates into the fatty acid bilayer. A spectrofluorometer is used to polarize UV light as a vertical excitation beam which passes through the dye-labeled membrane where the dye fluoresces. The beams perpendicular and horizontal to the excitation light are then collected and analyzed. Membrane structural properties are largely due to the packing of the fatty acids in the lipid bilayer that determines the membrane biophysical parameters. Staphylococcus aureus contains straight-chain (SCFAs) and branched-chain (BCFAs) fatty acids in the membrane and alters the proportion of membrane fluidizing BCFAs and stabilizing SCFAs as a response to a variety of stresses. Herein, we describe a method for determination of membrane fluidity in S. aureus using diphenylhexatriene, one of the most used fluorescent dyes for this purpose. Key words Staphylococcus aureus, Membrane fluidity, Fluorescence, Polarization, Anisotropy

1

Introduction Fatty acids in the membrane phospholipid bilayer largely determine membrane biophysical properties [1]. Gram-positive bacteria contain the cytoplasmic membrane as the only membranous structure. The membrane is a part of the cell envelope providing a physical barrier to the environment and is a site for energy (ATP) generation. In addition, the cell membrane is vital for cell division, virulence, maintenance of the electrochemical gradient, and transport of H2O, nutrients, ions, and proteins [2, 3]. Hence, cellular physiology is tightly connected with the optimal structure and function of the membrane which is a function of packing of fatty acids in the lipid bilayer [3]. S. aureus synthesizes straight-chain fatty acids (SCFAs) and branched-chain fatty acids (BCFAs) using fatty acid biosynthesis system II and alters the balance of membrane fluidizing BCFAs and SCFAs that increase membrane viscosity under different environmental conditions such as changes in temperature, pressure, ion

Kelly C. Rice (ed.), Staphylococcus aureus: Methods and Protocols, Methods in Molecular Biology, vol. 2341, https://doi.org/10.1007/978-1-0716-1550-8_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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concentrations, pH, nutrient availability and challenge with antimicrobials. Staphylococci lack fatty acid desaturase enzymes that convert SCFAs to straight-chain unsaturated fatty acids (SCUFAs), which are important membrane components in the cells of animals and plants, Gram-negative bacteria, and many Gram-positive bacteria including Streptococcus species. However, S. aureus can take SCUFAs from the growth environment and incorporate them into phospholipids [4, 5]. Additionally, many S. aureus strains produce a characteristic membrane component, staphyloxanthin, which is a triterpenoid pigment that rigidifies the membrane [5–7]. Fluorescence polarization methods are valuable for measuring bacterial cytoplasmic membrane fluidity [3]. A spectrofluorometer equipped with polarizers and photomultipliers is used to measure membrane fluidity [3, 8, 9]. A vertically polarized UV light excitation beam is passed through the sample in a cuvette and the emitted polarized light is collected in horizontal and vertical directions. Diphenylhexatriene (DPH) is the most commonly used fluorescent probe to label membranes [3]. DPH is a symmetrical and rod-like trans-polyene that intercalates in the hydrophobic fatty acid chains in the membrane and rotates in the lipid lattice at various rates based on the compactness of the membrane [3, 10]. The degree of membrane polarization (P) is calculated as the ratio of (IVV IVHG) to (IVV + IVHG) where IVV and IVH are emitted light intensities in the vertical and horizontal directions relative to the excitation light [3, 9]. G is the grating factor of the instrument referring to the sensitivities of the in-built photomultipliers [3, 11, 12]. Since there is a possibility of a second perpendicular emission plane oriented along the axis of propagation, another term, anisotropy (A), is preferred to accurately represent membrane fluidity. Anisotropy values account for the contribution of all angles of rotational freedom of the dye [13]. A is defined as the ratio of (IVV IVHG) to (IVV + 2IVHG) and is equal to 2P/3 P [3, 13]. The P and A values are inversely proportional to membrane fluidity [9].

2

Materials 1. 10 mM 1,6-Diphenyl-1,3,5-hexatriene (DPH) stock solution: Weigh 12 mg of DPH powder in low light (light off). Dissolve in 5 mL tetrahydrofuran (THF) in a clean glass tube covered with aluminum foil. Vortex 10 min in low light (see Notes 1 and 2). 2. 100 μM DPH working solution: Mix 20 μL of the DPH stock solution in 1980 μL of 1 PBS in an aluminum foil wrapped tube. Screw the cap tightly on the tube tightly and vortex thoroughly for 10 min. Evaporate THF by flushing with nitrogen gas for 10 min. Use a freshly prepared working solution for each experiment.

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3. 1 Phosphate buffered saline (1 PBS): Add approximately 800 mL H2O in a beaker containing a magnetic stir bar. Add the following components and gently mix on a magnetic plate stirrer until fully dissolved: 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, and 0.24 g of KH2PO4. Adjust pH to 7.4 (see Note 3). Transfer into a 1 L graduated cylinder and add H2O to a total volume of 1 L. Dispense into smaller volumes (100 mL in 250 mL screw cap bottles), autoclave and store at room temperature. 4. Nitrogen gas supply (see Note 4). 5. Tryptic Soy Broth (TSB): Add 1.5 g TSB powder to 50 mL H2O in a 250 mL flask. Mix until dissolved. Use a loose cap on the flask and autoclave with moist-heat (liquid) cycle at 121  C and 15 psi pressure for 15 min. 6. 1 M Tris–HCl, pH 7.6: Weigh 12.1 g of Tris-base in about 80 mL H2O, mix to dissolve the powder, and adjust pH using concentrated HCl (see Note 5). Transfer to a graduated cylinder and add H2O to achieve a final volume of 100 mL. Autoclave and store at room temperature. 7. 50 mM Tris–HCl, pH 7.6: Add 5 mL of 1 M Tris–HCl, pH 7.6–95 mL H2O. 8. Protoplast buffer: 20% (w/v) sucrose, 50 mM Tris–HCl, 145 mM NaCl, pH 7.6: Add 5 mL of 1 M Tris–HCl, pH 7.6 in approximately 70 mL distilled H2O in a 250 mL beaker containing a magnetic stir bar. Add the following components and gently mix on a magnetic plate stirrer until fully dissolved: 0.85 g NaCl, 20 g sucrose. Transfer the solution to a graduated cylinder and bring the volume to 100 mL with H2O. Filter sterilize using a 0.22 μm filter and store at 4  C until use. 9. Lysostaphin stock solution: 10 mg/mL lysostaphin in H2O. Store solution at 20  C. 10. DNase I stock solution: 2000 U/mL in H2O (see Note 6). 11. 14.3 M β-mercaptoethanol, 98% purity (see Note 7). 12. 50 mM β-mercaptoethanol: Dissolve 3.5 μL of 14.3 M β-mercaptoethanol in 1 mL H2O by mixing and inverting. Store at 4  C. 13. Bovine serum albumin (BSA) stock solution: Dissolve BSA in H2O to a final concentration of 1 mg/mL. 14. Shaking incubator. 15. 96-well plates for absorbance measurement, with flat clear bottom. 16. Spectrophotometer. 17. Plastic cuvettes (1 cm path length).

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Fig. 1 Spectrofluorometer. 1, High intensity xenon lamp; 2, Excitation monochromator; 3, Sample compartment; 4, Emission monochromator; 5, Detector

18. 50 mL Falcon tubes. 19. Water bath. 20. Benchtop centrifuge. 21. Refrigerated microcentrifuge. 22. Sonicator (see Note 8). 23. Ultracentrifuge. 24. 2 mL volume ultracentrifuge tubes. 25. Microplate reader. 26. Quartz cuvettes. 27. Spectrofluorometer (Fig. 1) (see Notes 9 and 10).

3

Methods

3.1 Sample Preparation

3.1.1 Whole Cell Suspension (Modified from [14, 15])

Since Gram-positive bacterial cells contain membrane as the only membranous structure, whole cell suspensions have been used to determine membrane fluidity with reproducible results [14, 15]. Protoplasts [16] can also be prepared and used for addressing research questions in which the results may be affected by the presence of the cell wall. Additionally, membranes themselves can be isolated and analyzed [17]. Isolated membrane samples may be the best for continuous analysis of membrane fluidity using a spectrofluorometer equipped with temperature adjustable cuvette holder. Therefore, all three sample preparation methods are outlined below. 1. Grow S. aureus cultures at 37  C and 250 rpm in 50 mL TSB in a 250 mL flask (1:5 volume to flask ratio) to optical density at 600 nm (OD600) ~1.0 (see Note 11).

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2. Transfer culture volumes to 50 mL Falcon tubes, and harvest cells by centrifugation (2500  g for 3 min). 3. Wash cell pellets twice with 1 PBS. 4. Resuspend cell pellets in 1 PBS to final OD600 of 0.45. 3.1.2 Protoplast Preparation (Modified from [16])

1. Grow S. aureus cultures at 37  C and 250 rpm in 50 mL TSB in a 250 mL flask to an OD600 ~1.0 (see Note 11). 2. Transfer culture volumes to 50 mL Falcon tubes, and harvest cells by centrifugation (5000  g for 15 min). 3. Resuspend cell pellets each in 1 mL protoplast buffer. 4. Digest the bacterial cell wall in each sample by adding 3 μL of lysostaphin stock (30 μg/mL final concentration) and 1 μL of DNase I stock (2 U/mL final concentration) to each 1 mL cell suspension (see Note 12). Mix gently and incubate for 1 h in a 37  C water bath. 5. Collect the protoplasts by centrifugation (10,000  g for 15 min at 4  C) and resuspend in fresh protoplast buffer.

3.1.3 Cell Membrane Extraction (Protocol Modified from [17])

1. Grow S. aureus cultures at 37  C and 250 rpm in 200 mL TSB in a 1 L flask to an OD600 ~1.0 (see Note 11). 2. Transfer culture volumes to 50 mL Falcon tubes, and harvest cells by centrifugation (5000  g for 15 min). 3. Wash the cell pellets with 1 PBS. 4. Resuspend each cell pellet in 2 mL 50 mM Tris–HCl pH 7.6. Add 10 μL of lysostaphin stock (50 μg/mL final concentration) and 6 μL of DNase I stock (6 U/mL final concentration) to each cell suspension (see Note 12). 5. Incubate cell suspensions for 30 min at 4  C. 6. Sonicate the protoplast/cell lysates for five cycles (30 s sonication followed by at least 1 min of incubation on ice). 7. Centrifuge all tubes at 15,000  g at 4  C for 15 min. 8. Transfer lysates from step 7 to ultracentrifuge tubes, and centrifuge at 50,000  g for 60 min at 4  C. 9. Resuspend the pellets (containing cell membranes) from step 8 in 200 μL 50 mM Tris–HCl, pH 7.6 containing 1 mM β-mercaptoethanol (add 4 μL of 50 mM β-mercaptoethanol stock per 200 μL volume of 50 mM Tris–HCl, pH 7.6). 10. Determine protein concentration by Bradford method [18]. Prepare BSA standards (1000, 500, 250, 100, 50, 25, 10 μg/mL final concentrations) by diluting the BSA stock solution in 50 mM Tris–HCl, pH 7.6 containing 1 mM β-mercaptoethanol. Add 200 μL Bradford reagent to each well of a 96-well plate. Add 10 μL of each standard and sample

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to duplicate wells, mix by pipetting, and incubate for 5 min at room temperature before reading the absorbance at 595 nm (A595) using a microplate reader. Include duplicate blank wells without BSA but containing 10 μL of 50 mM Tris–HCl, pH 7.6 containing 1 mM β-mercaptoethanol. Prepare a standard curve using the BSA standards and use to calculate the protein concentrations in the membrane samples. 11. Use 70–100 μg/mL equivalent membrane preparation per sample for anisotropy measurement (see Note 13). 3.2

DPH Treatment

Perform all steps involving DPH in low light (turn off lights). 1. Transfer a volume of each sample prepared in Subheading 3.1 (whole cells, protoplasts or cell membranes) each into a microcentrifuge tube, based on the volume of cuvette to be used for the spectrofluorometer. 2. Add 100 μM DPH working solution to final concentration of 2 μM to each tube (for example, add 20 μL of the DPH working solution to 980 μL of sample), and mix well (see Note 14). 3. Prepare a control for each sample that does not contain DPH. This will be used as a blank for each sample in step 9 of Subheading 3.3 below. 4. Wrap each tube with aluminum foil and incubate the tubes at 30  C in a H2O bath for 1 h (see Note 14).

3.3

Fluorometry

There are variety of spectrofluorometers available on the market. As an example of how to perform fluorometry for this experiment, we describe the use of a PTIModel Quanta Master-4 Scanning Spectrofluorometer (HORIBA) with FelixGX—4.2.2 software. Position the Excitation and Emission Lens firmly on inside wall of the sample compartment as prescribed and close the upper lid carefully. Switch the powerpads in a sequence as described and wait at least 15 min before setting up the program to measure the anisotropy values of the samples (see Notes 9 and 10). Open the FelixGX software and proceed as described below: 1. Click Setup > Acquisition type: Time-based. A new window named Setup: Acquisition will open. 2. Check on the following settings: Excitation, 360 nm; Emission, 430 nm; Excitation slit width, 5 nm, Emission slit width, 5 nm; Points/sec, 1; Duration, 60. Click Accept to proceed to the Setup window. 3. Click Setup > Polarizers > check Use Polarizers > click on G-Factor > click Accept to come back on the Acquisition window (Fig. 2). 4. Acquisition window > Traces > click Add on the lower panel to get a row to put information about the G-Factor. Double click

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Fig. 2 FelixGX software showing Setup and Setup:Acquisition windows

the box under Name > type G-Factor > select Source 1 as HH light component > select G-Factor under Function > select Source 2 as HV light component > uncheck Graph 1 and check Graph 2 > click Accept to get into the Setup window. 5. Click Start on the right side of Setup button to scan grating of the instrument (G-Factor, calibration of instrument) (Fig. 3). 6. After G-factor scan is done, right-click on the G-Factor trace in the list of instrument operation information panel (top left of the Setup window). Click Create Lookup Table > Name: G-factor > Type: G-Factor > OK to get on the Setup: Acquisition window (Fig. 4). 7. Acquisition > click on Polarization/Anisotropy > Use Lookup Table to get a new window > select G-Factor > OK > Accept > Accept. 8. Acquisition > click on Traces > click Add on the lower panel to get rows to put information about the Polarization and Anisotropy. Double click the box under Name > type Polarization > select Source 1 as HH light component > select Polarization under Function > select Source 2 as HV light component >

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Fig. 3 FelixGX software showing G-Factor scanned window

uncheck Graph 1 and check Graph 2. Similarly, put information for Anisotropy > then click Accept to get into the Setup window. The program will be ready to scan blank and samples (Fig. 5). 9. For each sample, add its corresponding blank (prepared in step 3 of Subheading 3.2) in a quartz cuvette, place in the cuvette holder of the sample compartment and close the lid. 10. Click Start next to the Setup button to start scanning the blank (Fig. 6). 11. Once the instrument has completed scanning the blank, remove the contents from the quartz cuvette, wash once with distilled H2O and once with ethanol, then dry the cuvette. Put the DPH-treated sample in the quartz cuvette, lace in the cuvette holder and click Start to scan the sample. The realtime scanning progress is observable on both the upper and lower panels of the Setup window. 12. After the sample scanning has been completed, click on the Graph-Grid button on the upper-left of the Setup window to

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Fig. 4 FelixGX software showing windows for how to link G-Factor to the polarization/anisotropy values

observe digital values of the scan over 60 s of DPH-membrane fluorescence (Fig. 7). 13. Save the output data as excel sheet/s. As shown in Fig. 7, various data points corresponding to the scanning time will appear as both Polarization and Anisotropy values. Average the data points for each sample and its corresponding blank. Subtract the blank from the polarization or anisotropy values to get the sample polarization or anisotropy values. 14. Representative data using this FP method to assess S. aureus membrane fluidity is depicted in Table 1, in which the anisotropy values of a pigmented S. aureus strain (Pig1) and its carotenoid-lacking strain (Pig1ΔcrtM) grown in TSB and human serum are shown.

4

Notes 1. DPH causes skin irritation, serious eye irritation and respiratory irritation. Store in dark and dry well-ventilated place. Keep container tightly closed. Work with this chemical in a wellventilated area and avoid dust and aerosol formation, wash skin thoroughly after handling, and wear appropriate personal

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Fig. 5 FelixGX software window showing how to add Polarization and Anisotropy parameters in fluorescence determination

protective equipment (PPE) such as gloves, goggles, and a mask. Prepare the stock solution in a screw cap glass tube covered with aluminum foil and store at 4  C. The shelf life of the stock solution is