Moonlighting Proteins E-Book

• Herausgeber: John Wiley & Sons
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 919

Veröffentlichungsjahr: 2017

Beschreibung

Moonlighting Proteins: Novel Virulence Factors in Bacterial Infections is a complete examination of the ways in which proteins with more than one unique biological action are able to serve as virulence factors in different bacteria. The book explores the pathogenicity of bacterial moonlighting proteins, demonstrating the plasticity of protein evolution as it relates to protein function and to bacterial communication. Highlighting the latest discoveries in the field, it details the approximately 70 known bacterial proteins with a moonlighting function related to a virulence phenomenon. Chapters describe the ways in which each moonlighting protein can function as such for a variety of bacterial pathogens and how individual bacteria can use more than one moonlighting protein as a virulence factor. The cutting-edge research contained here offers important insights into many topics, from bacterial colonization, virulence, and antibiotic resistance, to protein structure and the therapeutic potential of moonlighting proteins. Moonlighting Proteins: Novel Virulence Factors in Bacterial Infections will be of interest to researchers and graduate students in microbiology (specifically bacteriology), immunology, cell and molecular biology, biochemistry, pathology, and protein science.

Leseprobe

Table of Contents

Cover

Title Page

List of Contributors

Preface

About the Editor

Part I: Overview of Protein Moonlighting

1 What is Protein Moonlighting and Why is it Important?

1.1 What is Protein Moonlighting?

1.2 Why is Moonlighting Important?

1.3 Current questions

1.4 Conclusions

References

2 Exploring Structure–Function Relationships in Moonlighting Proteins

2.1 Introduction

2.2 Multiple Facets of Protein Function

2.3 The Protein Structure–Function Paradigm

2.4 Computational Approaches for Identifying Moonlighting Proteins

2.5 Classification of Moonlighting Proteins

2.6 Conclusions

References

Part II: Proteins Moonlighting in Prokarya

3 Overview of Protein Moonlighting in Bacterial Virulence

3.1 Introduction

3.2 The Meaning of Bacterial Virulence and Virulence Factors

3.3 Affinity as a Measure of the Biological Importance of Proteins

3.4 Moonlighting Bacterial Virulence Proteins

3.5 Bacterial Moonlighting Proteins Conclusively Shown to be Virulence Factors

3.6 Eukaryotic Moonlighting Proteins That Aid in Bacterial Virulence

3.7 Conclusions

References

4 Moonlighting Proteins as Cross‐Reactive Auto‐Antigens

4.1 Autoimmunity and Conservation

4.2 Immunogenicity of Conserved Proteins

4.3 HSP Co‐induction, Food, Microbiota, and T‐cell Regulation

4.4 The Contribution of Moonlighting Virulence Factors to Immunological Tolerance

References

Part III: Proteins Moonlighting in Bacterial Virulence

3.1 Chaperonins: A Family of Proteins with Widespread Virulence Properties

5 Chaperonin 60 Paralogs in

Mycobacterium tuberculosis

and Tubercle Formation

5.1 Introduction

5.2 Tuberculosis and the Tuberculoid Granuloma

5.3 Mycobacterial Factors Responsible for Granuloma Formation

5.4 Mycobacterium tuberculosis Chaperonin 60 Proteins, Macrophage Function, and Granuloma Formation

5.5 Conclusions

References

6

Legionella pneumophila

Chaperonin 60, an Extra‐ and Intra‐Cellular Moonlighting Virulence‐Related Factor

6.1 Background

6.2 HtpB is an Essential Chaperonin with Protein‐folding Activity

6.3 Experimental Approaches to Elucidate the Functional Mechanisms of HtpB

6.4 Secretion Mechanisms Potentially Responsible for Transporting HtpB to Extracytoplasmic Locations

6.5 Identifying Functionally Important Amino Acid Positions in HtpB

6.6 Functional Evolution of HtpB

6.7 Concluding Remarks

References

3.2 Peptidylprolyl Isomerases, Bacterial Virulence, and Targets for Therapy

7 An Overview of Peptidylprolyl Isomerases (PPIs) in Bacterial Virulence

7.1 Introduction

7.2 Proline and PPIs

7.3 Host PPIs and Responses to Bacteria and Bacterial Toxins

7.4 Bacterial PPIs as Virulence Factors

7.5 Other Bacterial PPIs Involved in Virulence

7.6 Conclusions

References

3.3 Glyceraldehyde 3‐Phosphate Dehydrogenase (GAPDH): A Multifunctional Virulence Factor

8 GAPDH: A Multifunctional Moonlighting Protein in Eukaryotes and Prokaryotes

8.1 Introduction

8.2 GAPDH Membrane Function and Bacterial Virulence

8.3 Role of Nitric Oxide in GAPDH Bacterial Virulence

8.4 GAPDH Control of Gene Expression and Bacterial Virulence

8.5 Discussion

Acknowledgements

References

9

Streptococcus pyogenes

GAPDH: A Cell‐Surface Major Virulence Determinant

9.1 Introduction and Early Discovery

9.2 GAS GAPDH: A Major Surface Protein with Multiple Binding Activities

9.3 AutoADP‐Ribosylation of SDH and Other Post‐Translational Modifications

9.4 Implications of the Binding of SDH to Mammalian Proteins for Cell Signaling and Virulence Mechanisms

9.5 Surface Export of SDH/GAPDH: A Cause or Effect?

9.6 SDH: The GAS Virulence Factor‐Regulating Virulence Factor

9.7 Concluding Remarks and Future Perspectives

References

10 Group B

Streptococcus

GAPDH and Immune Evasion

10.1 The Bacterium GBS

10.2 Neonates are More Susceptible to GBS Infection than Adults

10.3 IL‐10 Production Facilitates Bacterial Infection

10.4 GBS Glyceraldehyde‐3‐Phosphate Dehydrogenase Induces IL‐10 Production

10.5 Summary

References

11

Mycobacterium tuberculosis

Cell‐Surface GAPDH Functions as a Transferrin Receptor

11.1 Introduction

11.2 Iron Acquisition by Bacteria

11.3 Iron Acquisition by Intracellular Pathogens

11.4 Iron Acquisition by M. tb

11.5 Glyceraldehyde‐3‐Phosphate Dehydrogenase (GAPDH)

11.6 Macrophage GAPDH and Iron Uptake

11.7 Mycobacterial GAPDH and Iron Uptake

11.8 Conclusions and Future Perspectives

Acknowledgements

References

12 GAPDH and Probiotic Organisms

12.1 Introduction

12.2 Probiotics and Safety

12.3 Potential Risk of Probiotics

12.4 Plasminogen Binding and Enhancement of its Activation

12.5 GAPDH as an Adhesin

12.6 Binding Regions

12.7 Mechanisms of Secretion and Surface Localization

12.8 Other Functions

12.9 Conclusion

References

3.4 Cell‐Surface Enolase: A Complex Virulence Factor

13 Impact of Streptococcal Enolase in Virulence

13.1 Introduction

13.2 General Characteristics

13.3 Expression and Surface Exposition of Enolase

13.4 Streptococcal Enolase as Adhesion Cofactor

13.5 Enolase as Pro‐Fibrinolytic Cofactor

13.6 Streptococcal Enolase as Cariogenic Factor in Dental Disease

13.7 Conclusion

Acknowledgement

References

14 Streptococcal Enolase and Immune Evasion

14.1 Introduction

14.2 Localization and Crystal Structure

14.3 Multiple Binding Activities of α‐Enolase

14.4 Involvement of α‐Enolase in Gene Expression Regulation

14.5 Role of Anti‐α‐Enolase Antibodies in Host Immunity

14.6 α‐Enolase as Potential Therapeutic Target

14.7 Questions Concerning α‐Enolase

References

15

Borrelia burgdorferi

Enolase and Plasminogen Binding

15.1 Introduction to Lyme Disease

15.2 Life Cycle

15.3 Borrelia Virulence Factors

15.4 Plasminogen Binding by Bacteria

15.5 B. burgdorferi and Plasminogen Binding

15.6 Enolase

15.7 B. burgdorferi Enolase and Plasminogen Binding

15.8 Concluding Thoughts

Acknowledgements

References

3.5 Other Glycolytic Enzymes Acting as Virulence Factors

16 Triosephosphate Isomerase from 

Staphylococcus aureus

and Plasminogen Receptors on Microbial Pathogens

16.1 Introduction

16.2 Identification of Triosephosphate Isomerase on S. aureus as a Molecule that Binds to the Pathogenic Yeast C. neoformans

16.3 Binding of Triosephosphate Isomerase with Human Plasminogen

16.4 Plasminogen‐Binding Proteins on Trichosporon asahii

16.5 Plasminogen Receptors on C. neoformans

16.6 Conclusions

References

17 Moonlighting Functions of Bacterial Fructose 1,6‐Bisphosphate Aldolases

17.1 Introduction

17.2 Fructose 1,6‐bisphosphate Aldolase in Metabolism

17.3 Surface Localization of Streptococcal Fructose 1,6‐bisphosphate Aldolases

17.4 Pneumococcal FBA Adhesin Binds Flamingo Cadherin Receptor

17.5 FBA is Required for Optimal Meningococcal Adhesion to Human Cells

17.6 Mycobacterium tuberculosis FBA Binds Human Plasminogen

17.7 Other Examples of FBAs with Possible Roles in Pathogenesis

17.8 Conclusions

References

3.6 Other Metabolic Enzymes Functioning in Bacterial Virulence

18 Pyruvate Dehydrogenase Subunit B and Plasminogen Binding in

Mycoplasma

18.1 Introduction

18.2 Binding of Human Plasminogen to M. pneumoniae

18.3 Localization of PDHB on the Surface of M. pneumoniae Cells

18.4 Conclusions

References

3.7 Miscellaneous Bacterial Moonlighting Virulence Proteins

19 Unexpected Interactions of Leptospiral Ef‐Tu and Enolase

19.1 Leptospira –Host Interactions

19.2 Leptospira Ef‐Tu

19.3 Leptospira Enolase

19.4 Conclusions

References

20

Mycobacterium tuberculosis

Antigen 85 Family Proteins

20.1 Introduction

20.2 Identification of Antigen 85

20.3 Antigen 85 Family Proteins: Mycolyl Transferases

20.4 Antigen 85 Family Proteins: Matrix‐Binding Adhesins

20.5 Conclusion

Acknowledgement

References

3.8 Bacterial Moonlighting Proteins that Function as Cytokine Binders/Receptors

21 Miscellaneous IL‐1β‐Binding Proteins of

Aggregatibacter actinomycetemcomitans

21.1 Introduction

21.2 A. actinomycetemcomitans Biofilms Sequester IL‐1β

21.3 A. actinomycetemcomitans Cells Take in IL‐1β

21.4 The Potential Effects of IL‐1β on A. actinomycetemcomitans

21.5 Conclusions

References

3.9 Moonlighting Outside of the Box

22 Bacteriophage Moonlighting Proteins in the Control of Bacterial Pathogenicity

22.1 Introduction

22.2 Bacteriophage T4 I‐TevI Homing Endonuclease Functions as a Transcriptional Autorepressor

22.3 Capsid Psu Protein of Bacteriophage P4 Functions as a Rho Transcription Antiterminator

22.4 Bacteriophage Lytic Enzymes Moonlight as Structural Proteins

22.5 Moonlighting Bacteriophage Proteins De‐Repressing Phage‐Inducible Chromosomal Islands

22.6 dUTPase, a Metabolic Enzyme with a Moonlighting Signalling Role

22.7 Escherichia coli Thioredoxin Protein Moonlights with T7 DNA Polymerase for Enhanced T7 DNA Replication

22.8 Discussion

References

23 Viral Entry Glycoproteins and Viral Immune Evasion

23.1 Introduction

23.2 Enveloped Viral Entry

23.3 Moonlighting Activities of Viral Entry Glycoproteins

23.4 Viral Entry Proteins Moonlighting as Saboteurs of Cellular Pathways

23.5 Conclusions

References

Index

End User License Agreement

List of Tables

Chapter 02

Table 2.1 Proteins having distinct sites for different functions in the same domain.

Table 2.2 Proteins having distinct sites for different functions in different domains.

Table 2.3 Proteins using the same residues for different functions.

Table 2.4 Proteins using different residues in the same/overlapping site for different functions.

Table 2.5 Proteins using different folds for different functions.

Chapter 03

Table 3.1 Binding affinities of moonlighting proteins to various protein ligands.

Table 3.2 Binding affinities of moonlighting proteins to plasmino(gen).

Table 3.3 The distribution of bacterial moonlighting proteins into functional groups. Proteins in bold are those produced both by bacteria and eukaryotes.

Table 3.4 Some bacterial moonlighting proteins with (some of their) multiple moonlighting functions. Biological actions are not in any particular order, and not all the biological functions of these proteins are shown.

Table 3.5 Bacterial moonlighting proteins acting as bacterial adhesins.

Table 3.6 Bacterial moonlighting proteins functioning as evasins.

Chapter 07

Table 7.1 Bacteria with cell‐surface Mip‐like PPIs.

Chapter 08

Table 8.1 Membrane‐bound bacterial GAPDH as a virulence factor.

Table 8.2 GAPDH‐mediated transnitrosylation.

Chapter 09

Table 9.1 Post‐translational modifications of SDH and resulting moonlighting functions.

Table 9.2 SDH binding to mammalian proteins and resulting moonlighting functions.

Table 9.3 SDH‐mediated

S. pyogenes

virulence gene regulation (Jin

et al

. 2011).

Chapter 11

Table 11.1 Identified

M. tb

H37Rv transferrin‐binding proteins (Boradia

et al

. 2014).

Chapter 12

Table 12.1 Representative probiotic strains.

Table 12.2 Probiotics strains expressing GAPDH on cell surface and its targeting ligands.

Chapter 13

Table 13.1 Streptococcus enolases.

Table 13.2 Bacterial enolases.

Chapter 18

Table 18.1 Described glycolytic enzymes with multifunction in

Mycoplasma/Spiroplasma

.

Table 18.2 Results of reactivity of recombinant protein PDHB with sera (

n

 = 32) of patients with symptoms of community‐acquired pneumonia and confirmed

M. pneumoniae

‐specific antibodies in comparison with total antigen of

M. pneumoniae

(ELISA; secondary antibody: anti‐human whole Ig‐HRP).

Chapter 19

Table 19.1

Leptospira

proteins exhibiting moonlighting activities.

Chapter 23

Table 23.1 Viral moonlighting proteins and their interactions with the complement system.

Table 23.2 Role of various viral GPs in antibody evasion.

Table 23.3 Moonlighting activities of the HIV‐1 gp41 viral entry glycoprotein.

List of Illustrations

Chapter 02

Figure 2.1 Function annotations for the mouse protein, glucose‐6‐phosphate isomerase (Uniprot Accession no. P06745) from the Enzyme Commision (EC) number system and Gene Ontology.

Figure 2.2 From protein structure to function.

Figure 2.3 α‐Enolase. (a) Single chain of Enolase showing the enzyme active site in blue and the plasminogen‐binding site in red. (b) Enolase monomer displayed as surface. Different domains are colored in gray and orange (PDB:1W6T).

Figure 2.4 Albaflavenone monooxygenase. The monooxygenase and terpene synthase active sites are shown in blue and red respectively in the (a) cartoon and (b) surface representation of Albaflavenone monooxygenase (PDB: 3EL3).

Figure 2.5 Human MAPK1/ERK2. The MAPK1 active site is shown in blue and the DNA‐binding motif is highlighted in red. Different domains are shown in gray and orange (PDB:4G6N).

Figure 2.6 Malate synthase. The enzyme active site is shown in blue and the laminin‐ binding site is shown in red. Different domains are shown in different colors (PDB:2GQ3).

Figure 2.7 BirA. The catalytic site residues are shown in blue while the H‐T‐H motif involved in binding DNA (moonlighting function) is shown in red. Different domains are shown in different colors (PDB:1BIB).

Figure 2.8 Human MRDI. The active site residues are shown in blue while the residues implicated in controlling invasion (moonlighting function) is shown in red. Different domains are shown in different colors (PDB:4LDQ).

Figure 2.9 GAPDH. The catalytic site residue Cys149 (shown in red) is the residue known to be involved for both the canonical and moonlighting functions of

E. coli

GAPDH. The other catalytic residue His179 is shown in blue (PDB:1DC5).

Figure 2.10 Leukotriene A4 hydrolase. The LTA4 catalytic site residues Glu296 and Tyr383 are shown in blue. The catalytic site residue Glu271, involved in two separate functions in two different catalytic reactions is shown in red (PDB:2R59).

Figure 2.11 Phosphoglucose isomerase (PGI). Catalytic residues are shown as red sticks. Inhibition of enzymatic and AMF functions of PGI by the PGI inhibitor and mutational analysis of the catalytic residues have indicated overlapping regions of both functions in the human PGI (PDB:1IAT).

Figure 2.12 Aldolase. The enzyme active site is shown in blue and the actin‐binding site is shown in red (PDB:2PC4).

Figure 2.13 RfaH The Rfah CTD is colored in orange. In the closed form of RfaH (a), the CTD (α‐helix form) and NTD tightly interacts and works as a transcription factor (PDB:2OUG). The subsequent (or simultaneous) refolding of the CTD into a (b) β‐barrel transforms RfaH into a translation factor (PDB:2LCL).

Figure 2.14 Structural diversity v. functional diversity of CATH domain superfamilies represented in the moonlighting proteins studied in this chapter. Structural diversity is represented by the number of structural clusters (domains clustered at 5Å RMSD) in the superfamilies and the functional diversity is represented by the number of functional families identified in the superfamily.

Chapter 03

Figure 3.1 A schematic diagram showing the dynamic “timeline” of a typical bacterial infection and the requirements for the major virulence factors in the process.

Chapter 05

Figure 5.1 A cartoon view of the tuberculoid granuloma with cellular compositions deemed to be associated with bactericidal action or favoring bacterial persistence.

Chapter 06

Figure 6.1 Signaling cascades used by HtpB to trigger pseudohyphal growth in

S. cerevisiae

. Diagram showing the natural signaling pathways leading to expression of genes (

FLO11

being a main one) required for pseudohyphae formation. HtpB acts at the level of, or upstream from Ras2, signaling in a nitrogen‐ and glucose‐independent manner. Our data indicate that the Ste‐kinase and cAMP/PKA pathways are both critical to, and the only two pathways involved in, the induction of pseudohyphal growth by HtpB. Factors from top to bottom: Gpr1: G protein‐coupled receptor/transmembrane sensor; Gpa2: Gα subunit; Msb2: sensor/receptor; Sho1 and Opy2: sensors/G protein adaptors; Cdc24: guanine nucleotide exchange factor for Cdc42; Ras2: G protein global regulator; Cyr1: adenylate cyclase; Cdc42: Rho GTPase; Bcy1: regulatory subunit of the PKA complex; Tpk1/2/3: protein kinases A; Yak1: dual‐specificity tyrosine‐regulated kinase; Ste20: p21‐activated kinase; Ste11: MAPKKK; Ste50: adaptor/scaffold protein; Ste7: MAPKK; Kss1: MAPK of the filamentous growth pathway; Fus3: MAPK of the mating pathway; Flo8/Sok2/Ste12/Tec1/Phd1: transcription factors;

FLO11

: main hub of signal transduction. Notes: Upon binding cAMP, Bcy1 releases the three PKAs Tpk1/2/3, which then become active. Yak1 is active when non‐phosphorylated and it mediates low expression of Flo11 (thin arrows). Phosphorylation of Yak1 by Tpk1 inactivates it (gray arrow with circular head). Ste50 is known to specifically bind Ste11, but it could serve as the scaffold for the MAP kinase module (Ste11 + Ste7 + Kss1). Unregulated filamentous growth is detrimental for mating, so activated Fus3 acts as a repressor of Tec1 + Ste12 transcription factors (gray arrow with circular head). Active transcription from the

FLO11

promoter is depicted by the curved white arrow on the left of the

FLO11

gene.

Figure 6.2 HtpB‐induced filamentation and agar invasion (but not elongation and unipolar budding) requires the Ste‐kinase pathway in

S. cerevisiae

. Light micrographs of microcolonies of

S. cerevisiae

strain W303‐1b with the indicated deletions in genes encoding members of the Ste‐kinase (MAP‐kinase) pathway, and either containing pPP389::

htpB

(Nasrallah

et al

. 2011

b

) for expression of HtpB (+HtpB), or the empty vector control pPP389 (VC). All deletion mutants were grown on inducing galactose solid medium, as HtpB expression is controlled by a galactose‐inducible promoter. The

ste

mutants were created by allelic replacement with restriction fragments from plasmids pEL45 (

ste20

Δ::

URA3

) (Leberer

et al

. 1992), pSL1311 (

ste11

Δ::

URA3

), pSL1077 (

ste7

Δ::

URA3

), and pSL1094 (

ste12

Δ::

URA3

) (Stevenson

et al

. 1992). The restriction enzymes used were: pEL45

Xba

I/

Sal

I, pSL1077

Kpn

I/

Cla

I, pSL1094

Bam

HI/

Xho

I, and pSL1311

Sst

I/

Sph

I. The restriction fragments were transformed as linear DNA into strain W303‐1b, and colonies of deletion mutants were then selected on solid medium lacking uracil. Bars represent 12.5 µm.

Figure 6.3 HtpB induces cell elongation and unipolar budding in

S. cerevisiae

strain W303‐1b carrying the dominant negative

CDC42

Ala‐118

allele. Light micrographs of microcolonies (grown on inducing galactose solid medium) of

S. cerevisiae

strain W303‐1b bearing: (a) plasmid YCp(

CDC42

Ala‐118

) for expression of Cdc42p‐Ala118 (Ziman

et al

. 1991); or (b, c) plasmids pLM86 (Nasrallah

et al

. 2011

b

) and YCp(

CDC42

Ala‐118

) for simultaneous expression of HtpB and Cdc42p‐Ala118 (+HtpB in the presence of YCp(

CDC42

Ala‐118

)). Plasmid YCp was generously donated by H.‐U. Mösch (Philipps University, Germany). Yeast cells carrying the

CDC42

Ala‐118

allele grew large but did not elongate (a). Some large elongated cells are marked with the white arrow in (b), and a chain of elongated cells derived from a single large round cell is marked by the arrowheads in (c). Bars represent 10 µm in all panels.

Figure 6.4 HtpB‐induced filamentation and agar invasion (but not elongation and unipolar budding) also requires the cAMP/PKA signaling pathway. Light micrographs of microcolonies of haploid cells of

S. cerevisiae

strain BY4741 (

MAT

a

flo8::kan

r

his3‐11 leu2‐3,112 met15 ura3‐52

), which is a

flo8

Δ derivative of strain S288C (Brachmann

et al

. 1998). Strain BY4741 was either bearing: (a) plasmid pLM87 for expression of HtpB (Nasrallah

et al

. 2011

b

); (b) plasmid pHL135 for expression of Flo8p (Source: Liu

et al

. 1996); or (c) both plasmids pLM87 and pHL135 (HtpB and Flo8p). The elongated cells shown in (a) budded in a unipolar manner, but were not agar‐invasive. The

FLO8

‐complemented cells shown in (b) are aggregated and did not elongate or penetrate the agar. The cells shown in (c) had penetrated the agar (they were hyper‐invasive). Bars represent 10 µm.

Figure 6.5 HtpB‐induced pseudohyphal development in

S. cerevisiae

is dependent on Ras2p. Light micrographs of microcolonies grown on inducing galactose solid medium of a

ras2

Δ mutant strain (W303‐1b

ras2

deletion mutant) bearing the following plasmids: (a) pPP389::

htpB

(for expression of HtpB; Nasrallah

et al

. 2011

b

); (b) pRS313::

RAS2

(for expression of Ras2p) plus empty vector pPP389; and (c) pPP389::

htpB

and pRS313::

RAS2

(HtpB and Ras2p). To create the

ras2

Δ mutant,

RAS2

in strain W303‐1b was replaced with the allele

ras2

::

URA3

cut with

Eco

RI/

Hin

dIII from plasmid p

ras2

::

URA3

(Kataoka

et al

. 1984) and transformed as linear DNA. Colonies of

ras2

Δ mutant cells were selected on solid medium lacking uracil. For genetic complementation,

RAS2

was amplified by PCR from strain W303‐1b using primers

RAS2

full‐F (GTGGCCGTATCAATGGATC) and

RAS2

full‐R (GGGAAAGAGAAGCTTGTTATTC). The 1967 bp PCR amplification product was digested with

Xba

I, and cloned into the

Xba

I and

Eco

RV sites of the low‐copy number yeast plasmid pRS313 (Sikorski and Hieter 1989) to generate pRS313::

RAS2

. Note that complementation of

ras2

Δ cells with wild‐type

RAS2

alone did not result in formation of pseudohyphae (b). Bars represent 12.5 µm.

Figure 6.6 A Cpn60‐specific secretion mechanism exists in

B. pertussis

and is induced by the BvgAS two‐component regulatory system. Electron micrographs of ultrathin sections of specimens fixed with 4% freshly depolymerized paraformaldehyde, and subjected to immunogold labeling using a rabbit polyclonal antibody (Chong

et al

. 2009) that recognizes Cpn60, and a secondary goat anti‐rabbit gold conjugate (Sigma Immunochemicals). A wild‐type

B. pertussis

parent strain (Bp388 (wt)), a mutant with a defective Ptl type IV secretion system (Ptl –), and a mutant with a non‐functional BvgAS two‐component regulatory system (Bvg –) are shown. For size reference, the gold spheres that indicate the localization of Cpn60 epitopes are 10 nm in diameter.

Figure 6.7 Phylogenetic relationships between 48 chaperonins (belonging to 35 bacterial species) known to have moonlighting functions and (or) extra‐cytoplasmic locations. (a) Diagram of an unrooted maximum likelihood phylogenetic tree of the 48 chaperonins. Colors indicate phylum as per legend. Asterisk marks the highly divergent Cpn60.2 and Cpn60.3 of

Chlamydia trachomatis

and

Chlamydophila pneumoniae

, whereas the black arrowhead shows the positions of the Cpn60.1 of these two chlamydiales, which are more closely related to the Cpn60 of

Borrelia burgdorferi

,

Porphyromonas gingivalis

, and

Leptospira interrogans

(black branches between the two purple triangles). The open triangle marks the group formed by the three Cpn60s of

Mycoplasma gallisepticum

,

Mycoplasma genitalium

, and

Mycoplasma pneumoniae

, whereas the pound sign marks the position of the

Mycoplasma penetrans

Cpn60, which closely groups with the

Helicobacter pylori

’s Cpn60. Branch lengths are proportional to the number of amino acid substitutions per site (scale shown at the bottom of the diagram). (b) Diagram of part of the rooted tree of the same 48 chaperonins shown in (a). The part shown would be equivalent to the central part of the Proteobacteria branch (red bubble) depicted in (a), and includes HtpB (marked with the black arrow) and its closest moonlighting Cpn60s from the intracellular pathogens

Piscirickettsia salmonis

and

Francisella tularensis

. The tip labels include the amino acid sequence identification number (Gis for the Pfam database) followed by the bacterial taxonomic name. Branch lengths are proportional to the number of substitutions per site (scale given at the top of the panel). Bootstrap values (%) based on 100 replications are given for every branch.

Chapter 08

Figure 8.1 Functional diversity of GAPDH.

Figure 8.2 Membrane‐associated GAPDH: structure and function.

Figure 8.3 GAPDH and nitric oxide: multipotential effects.

Figure 8.4 GAPDH and gene regulation.

Figure 8.5 Mechanisms of genomic potentiation.

Chapter 09

Figure 9.1 Schematic diagram showing the role of SDH in the

S. pyogenes

pathogenesis. (a) Various stages of

S. pyogenes

infection. (b) During infection, as a successful pathogen

S. pyogenes

adheres to and invades host cells. Subsequently, it hides and proliferates within the host tissues by evading host innate immune responses and maintains its abundance in a local microenvironment.

S. pyogenes

also causes apoptosis of host cells. In all these different stages of the infection, SDH participates in almost all stages by binding through different receptor ligands as illustrated. (c) The SDH surface export is also important to maintain GAS virulence as, by retaining SDH within cytoplasm, 128 genes, including 25 major virulence genes, are downregulated. SDH is therefore a quintessential important virulence regulator. The surface export of the cytoplasmic SDH and its role in the

S. pyogenes

virulence regulation may be mediated via several post‐translational modifications including reported phosphorylation, ADP‐ribosylation, S‐guanylation, and ubiquitination. The mechanism underlying this possible and predicted regulation is however unknown.

Chapter 10

Figure 10.1 Group B

Streptococcus

GAPDH modulates the neonatal inflammatory response. (a) GBS cocci growing in chains may spontaneously lyse (dashed red cocci) and the released cytosolic GAPDH (green dot) can then re‐associate with living bacteria. Free or GBS‐bound GAPDH can interact with (b) B cells or with (c) macrophages. Upon interaction with B cells, GAPDH (d) triggers IL‐10 production and (e) impairs neutrophil recruitment. (f) GAPDH can also interact with macrophages to induce their apoptosis.

Chapter 11

Figure 11.1

M. tuberculosis

expresses cell‐surface transferrin‐binding proteins. (a) FACS analysis indicates specific binding of Tf‐Alexa647 which is competitively inhibited in the presence of 200X unlabeled transferrin. (b)Transferrin conjugated gold particles localize on the surface of intact

M. tb

H37Ra. (c) No surface labeling is observed in controls (Streptavidin‐gold conjugate). Scale bar represents 0.2 µm.

Figure 11.2 Transferrin and GAPDH co‐localize at the cell surface. (b) Co‐localization of GAPDH and transferrin by immunogold labeling transmission electron microscopy of

M. tb

H37Ra strain overexpressing GAPDH. GAPDH was detected using 1:100 polyclonal rabbit α‐GAPDH, followed by secondary antibody conjugated with 5 nm gold particles. Cells were simultaneously labeled with transferrin‐20 nm gold conjugate. Inset: magnified image of highlighted area showing co‐localization of GAPDH and transferrin. Arrows indicate 5 nm particles (GAPDH); arrowheads indicate the presence of transferrin‐gold. (a) Cells labeled with an equivalent amount of rabbit pre‐immune sera (controls). Scale bar represents 100 nm.

Figure 11.3 Transferrin‐iron uptake is mediated by the internalization of transferrin. Presence of transferrin gold particles within the cytoplasm of

M. tb

H37Ra cells after incubation at 37°C for 1 hr. Scale bar represents 100 nm.

Figure 11.4 Mechanisms for the uptake of transferrin iron and Heme by

M. tb

. 1, 2. Iron depletion stimulates the recruitment of

M. tb

GAPDH to the cell surface. 3, 4. Surface GAPDH captures holo‐transferrin and the entire complex is internalized within the bacilli. 5, Iron is released from transferrin; its subsequent fate regarding degradation or recycling out of the bacterium is not known. 6, Carboxymycobactin acquires transferrin iron, which is then delivered to the bacilli via mycobactin. 7, Carboxymycobactin directly utilizes the high affinity importer Irt AB to deliver transferrin iron. 8, Heme utilization by

M. tb

.

Chapter 12

Figure 12.1 (a) A representative structure of human intestinal mucus. A dashed square indicates expression of A‐type blood group antigen. (b) Chemical structure of three kinds of human ABO blood group antigens conjugated‐BSA probes. GalNAc: N‐acetylgalactosamine; Gal: galactose; Fuc: fucose; GlcNAc: N‐acetylglucosamine; NeuAc: N‐acetylneuraminic acid (sialic acid); Ser/Thr: serine or threonine.

Figure 12.2 The three‐dimensional structure and amino acid sequences of GAPDH of

L. plantarum

LA 318. The underline indicates CXXXC motif.

Figure 12.3 Overview diagram of GAPDH in probiotics.

Chapter 13

Figure 13.1 Scheme of moonlighting functions of streptococcal enolase in four categories: interaction with fibrinolysis; stress response; intracellular activities; and function as adhesion cofactor. Enolase structure

Figure 13.2 (a) Electron microscopic co‐localization of enolase and GAPDH on the surface of

Streptococcus canis

G2 by immune‐gold labeling performed by Manfred Rohde, Helmholtz Center for Infection Research (HZI), Braunschweig. (b) FESEM‐visualization at 50,000× magnification and at 100,000× magnification. Arrows point to protein A gold particle (15 nm) detecting GAPDH and arrow heads mark protein A gold (10 nm) detecting enolase.

Chapter 14

Figure 14.1 Multimeric crystal structure of

S. pneumoniae

and human α‐enolase.

S. pneumoniae

α‐enolase (PDBID: 1w6t) forms an octamer and the human version (PDBID: 3B97) is a dimer. This figure was produced using Yorodumi (run by PDBj) with Jmol, available at http://pdbj.org/emnavi/viewtop.php.

Figure 14.2 Phylogenetic tree of

Streptococcus

α‐enolase. The tree is calculated and drawn by “Average Distance using BLOSUM62” on Jalview 2.8.2. The amino acid sequences were obtained from each genome referential strains.

Figure 14.3 Multiple roles of α‐enolase in

Streptococcus

. α‐enolase has been shown to play various roles in bacterial cell cytoplasm and on the cell surface, as well as in culture supernatant. In the cytoplasm, α‐enolase functions as a glycolytic enzyme in energy metabolism and is also involved in gene regulation. α‐enolase interrupts the host immune system by binding host factors, such as plasminogen and C4b‐binding protein. α‐enolase is an immunogenic protein and antibodies cross‐react with human enolase. In addition, α‐enolase interacts with neutrophils and induces formation of NETs.

Chapter 15

Figure 15.1 Alignment of enolase protein sequences from human (three isoforms),

Staphylococcus aureus

,

Escherichia coli

, and

B. burgdorferi

. Identical amino acids are in red; similar amino acids are in blue font. A consensus line underneath the alignment indicates identity (*) or similarity (.). The box at the top of the alignment indicates a hydrophobic domain thought to play a role in membrane association (Pancholi 2001). The box at the bottom of the alignment indicates the catalytic site of the enzyme. Yellow highlighted text indicates an internal plasminogen‐binding motif (Noguiera

et al

. 2012). Bold black text indicates cross‐reactive epitopes associated with autoimmunity in cancer (Adamus

et al

. 1998).

Chapter 16

Figure 16.1 Protein–protein and protein–carbohydrate interactions between microbes and human hosts.

Chapter 18

Figure 18.1 Binding of human plasminogen to

M. pneumoniae

. (a) Immunofluorescence of fixed

M. pneumoniae

cells after incubation with Atto488‐labeled plasminogen (50 µg mL

–1

). Antiserum to Triton X‐insoluble proteins detected with TRITC‐labeled secondary antibody is used as control. Bar: 10 µm. (b) Ligand immunoblotting assay using SDS‐PAGE‐separated total proteins of

M. pneumoniae

incubated with plasminogen (15 µg mL

–1

) detected with rabbit anti‐plasminogen.

Figure 18.2 (a) Time‐dependent binding of human plasminogen (2.5 µg mL

–1

) after incubation with

M. pneumoniae

cells. (b) Detection of human plasminogen in immunoblotting using rabbit anti‐plasminogen.

Figure 18.3 Degradation of human fibrinogen (10 µg mL

–1

) in the presence of plasminogen (Plg; 10 µg/mL

–1

), recombinant protein PDHB (20 µg mL

–1

) and activator human uPA (4 ng mL

–1

). Fibrinogen was separated, blotted, and detected by rabbit anti‐fibrinogen.

Figure 18.4 Time‐dependent quantitative occurrence of PDHB in membrane and cytosolic fraction of total proteins of

M. pneumoniae

M129 under different conditions of incubation as measured by ELISA. Dilutions of recombinant protein PDHB of known concentration were used as standard.

Figure 18.5 Occurrence of subunit B of PDH complex in membrane and cytosolic fraction of total proteins of mutants B170 (lacking adherence‐related proteins P40 and P90) and Iv22a (lacking main P1 adhesin) in comparison to type strain M129 of

M. pneumoniae

. Detection of cytosolic enzyme enolase (Eno) was carried out as control.

Figure 18.6 ELISA reactivity of total proteins of

M. pneumoniae

and recombinant protein PDHB (10 µg mL

–1

each) with guinea pig antisera obtained after intranasal (i.n.) infection of animals, after subcutaneous immunization with total antigen (s.c. wM.p.), with fractions of Triton X‐100 insoluble proteins (s.c. TX ins) and cytosolic proteins (s.c. cyt) of

M. pneumoniae

.

Chapter 20

Figure 20.1 An illustrated summary of the multiple roles of Ag85. Ag85 (gray surface) is positioned relative to the mycomembrane (gray mesh) so the trehalose (black surfaces) binding sites are accessible from the membrane surface. The bound locations of host protein domains (white surfaces) are based on experimental models.

Chapter 21

Figure 21.1 Amino acid sequence of a novel bacterial interleukin receptor I (BilRI), which is an outer membrane (signal sequence underlined) lipoprotein (lipidation site marked with *). The protein contained a triplicate repetitive 40 amino acid long sequence after the first 50 amino acids (including the signal sequence).

Figure 21.2 Clustal O sequence alignment of the

A. actinomycetemcomitans

BilRI amino acid sequence with three different variants of

H. ducreyi

fibrinogen binder A (FgbA).

Figure 21.3 Viable

A. actinomycetemcomitans

cells internalize IL‐1β both (a) in biofilm and (b) as planktonic cells. Pre‐grown

A. actinomycetemcomitans

biofilms were co‐cultured with an organotypic gingival mucosa, and the localization of IL‐1β was investigated using immuno‐electron microscopy. IL‐1β was detected both attached to the inner membrane (IM) and in the outer edges of nucleoids (N).

Chapter 22

Figure 22.1 The homing mechanism for mobile group I introns. The endonuclease and flanking introns are transcribed, splicing occurs, and the homing endonuclease is translated as a free‐standing protein. The endonuclease activity will cleave an uninterrupted host gene at the corresponding cleavage site, forming a double‐strand break (DSB) close to the intron insertion site. Repair by homology‐driven strand invasion, recombination, and replication using the intron‐containing allele leads to incorporation of the intron and associated endonuclease into the gene.

Figure 22.2 A cartoon depicting the two separate functions of homing endonuclease I‐TevI. (a) This section shows the homing function of I‐TevI in which the helix‐turn‐helix interacts with the AT‐rich section of a homing site, leading to the zinc finger directing the catalytic domain 23–25 nucleotides upstream of the intron integration site (Intron Int Site). The catalytic domain cleaves as the CXXXG site and this cleavage forms the DSB necessary for intron homing and incorporation. (b) This section depicts the autorepressor function of the I‐TevI. In this case the helix‐turn‐helix interacts with an AT‐rich site displaying 15 bp identity to the homing AT‐rich site, but upstream of an operator site. This operator AT‐rich sequence may overlap with the T4 late promotor. Binding of the helix‐turn‐helix to the operator site directs the catalytic domain to interact with a CXXXG site 9 nt closer to the DNA‐binding site. Cleavage at this site is reduced

c

. 100‐fold compared to at the homing site, and this reduction in catalytic activity silences transcription.

Figure 22.3 (a) The crystal structure of the Psu monomer with seven α‐helices and dimerization interface. The N‐terminal, α1, and α2 helices form a tight coiled‐coil structure termed the “CC‐stem,” surrounded by the three C‐terminal helices (α5–α7) known as the CT‐belt. The central region, shown here as the dimerization interface and consisting of helices α3 and α4, forms a knotted structure with another monomer. CC‐stem in cyan, CT‐belt in green and central region in magenta. (b) The dimeric structure shown here is formed when the two Psu monomers “knot,” and this knotted region (as indicated) acts as a hinge allowing the other regions to swing inward and outward. This dimer formation allows Psu to interact with and bind the Rho hexamer. (PDB code: 3RX6). The figure shows the dimer with domain colour differences. (c) A top view of the closed, hexameric structure of Rho bound to RNA (PDB code: 3ICE). The two Psu‐binding regions (PBR1 and PBR2) are indicated by arrows along with the central region containing the RNA.

Figure 22.4 (a) A cartoon representation of a Psu dimer (PDB: 3RX6) interacting with a Rho hexamer (PDB:3ICE). The Psu dimer occupies diagonally opposite intersubunit niches of the hexamer, near Rho ATP‐binding sites. This interaction positions the Psu dimer over the central channel of the Rho hexamer, which is predicted to mechanically impede the translocase activity of the Rho. (b) A cartoon showing the action of Psu as a transcription antiterminator. The initial figure shows the Rho hexamer on the nascent RNA of the elongation complex (EC), which will lead to the termination of transcription once Rho disassociates the EC from the RNA and template DNA. The second image represents the antitermination mechanism, in which Psu dimer blocks the central channel of the Rho, preventing the translocation of nascent RNA through the Rho and thus preventing the disassociation of the EC from the RNA and template DNA. The Psu dimers here are depicted by surface representation.

Figure 22.5 SaPI induction is mediated by a helper phage‐encoded protein. The helper phage and the SaPI reside stably integrated into the bacteria chromosome due to the activity of their respective encoded master repressors, CI for the phage and Stl for the SaPI. When the SOS response is activated the repressor of the phage CI is degraded, allowing the expression of the lytic genes of the phage and initiating its replication. The interaction of a phage‐encoded protein (Inducer) with the Stl repressor of the SaPI allows the de‐repression of the main SaPI promoters and the activation of the island.

Chapter 23

Figure 23.1 Illustration of a typical viral fusion event catalyzed by viral envelope and entry glycoproteins with moonlighting activities.

Figure 23.2 Shed, soluble, and virion attached Ebola virus glycoproteins.

Figure 23.3 A schematic diagram placing the moonlighting activities of the HIV‐1 gp41 viral entry glycoprotein into a structural context.

Guide

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Table of Contents

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Moonlighting Proteins: Novel Virulence Factors in Bacterial Infections

Edited by Brian Henderson

Division of Infection and Immunity, University College London, London, UK

Copyright © 2017 by John Wiley & Sons, Inc. All rights reserved

Published by John Wiley & Sons, Inc., Hoboken, New Jersey

Published simultaneously in Canada

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Library of Congress Cataloging‐in‐Publication Data

Names: Henderson, Brian (Professor) editor.Title: Moonlighting proteins : novel virulence factors in bacterial infections / Edited by Brian Henderson.Description: Hoboken, New Jersey : John Wiley & Sons, Inc., [2017] | Includes bibliographical references and index.Identifiers: LCCN 2016049475 (print) | LCCN 2016051302 (ebook) | ISBN 9781118951118 (cloth) | ISBN 9781118951125 (pdf) | ISBN 9781118951132 (epub)Subjects: | MESH: Bacterial Proteins–physiology | Virulence Factors | Bacterial InfectionsClassification: LCC RA644.B32 (print) | LCC RA644.B32 (ebook) | NLM QW 52 | DDC 614.5/7–dc23LC record available at https://lccn.loc.gov/2016049475

Cover Image: © SEBASTIAN KAULITZKI/SCIENCE PHOTO LIBRARY/Getty ImagesCover Design: Wiley

List of Contributors

Angela Barbosa Laboratório de Bacteriologia, Instituto Butantan, São Paulo, Brazil.

Simone Bergmann Department of Infection Biology, Institute of Microbiology, Technische Universität Braunschweig, Braunschweig, Germany.

Vishant M. Boradia Department of Biotechnology, National Institute of Pharmaceutical Education and Research (NIPER), SAS Nagar, Punjab, India; Present address: Department of Microbiology and Immunology, University of Maryland School of Medicine, Baltimore, Maryland, USA.

Janine Z. Bowring Institute of Infection, Immunity and Inflammation, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK.

Catherine A. Brissette Department of Microbiology, University of North Dakota School of Medicine and Health Sciences, Grand Forks, North Dakota, USA.

Yung‐Fu Chang Department of Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA.

Jonathan D. Cook Department of Laboratory Medicine and Pathobiology, Faculty of Medicine, University of Toronto, Toronto, Canada.

Sayoni Das Institute of Structural and Molecular Biology, University College London, Gower Street, London, UK.

Roger Dumke Technical University Dresden, Dresden, Germany.

Paula Ferreira ICBAS, Instituto de Ciências Biomédicas de Abel Salazar, Universidade do Porto, Porto, Portugal; Instituto de Investigação e Inovação em Saúde, Universidade do Porto, Porto, Portugal

Kathleen Friedrich Technical University Dresden, Dresden, Germany.

Marcus Fulde Department of Infectious Diseases, Institute for Microbiology, University of Veterinary Medicine Hannover, Hannover, Germany; Present address: Centre for Infection Medicine, Institute of Microbiology and Epizootics, Freie Universität Berlin, Berlin, Germany.

Rafael A. Garduño Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada; Present address: Canadian Food Inspection Agency, Dartmouth Laboratory, Dartmouth, Nova Scotia, Canada.

Anne Gründel Technical University Dresden, Dresden, Germany.

Brian Henderson Department of Microbial Diseases, UCL‐Eastman Dental Institute, University College London, UK.

Tomoe Ichikawa Department of Microbial Science and Host Defense, Meiji Pharmaceutical University, Tokyo, Japan.

Riikka Ihalin Department of Biochemistry, University of Turku, Turku, Finland.

Reiko Ikeda Department of Microbial Science and Host Defense, Meiji Pharmaceutical University, Tokyo, Japan.

Chaaya Iyengar Raje Department of Biotechnology, National Institute of Pharmaceutical Education and Research (NIPER), SAS Nagar, Punjab, India.

Enno Jacobs Technical University Dresden, Dresden, Germany.

Constance J. Jeffery Department of Biological Sciences, University of Illinois at Chicago, Chicago, Illinois, USA.

Shigetada Kawabata Department of Oral and Molecular Microbiology, Osaka University Graduate School of Dentistry, Suita, Osaka, Japan.

Ishita Khan Department of Computer Science, Purdue University, North University Street, West Lafayette, Indiana, USA.

Daisuke Kihara Department of Biological Sciences, Purdue University, Martin Jischke Drive, West Lafayette, Indiana, USA.

Hideki Kinoshita Department of Food Management, School of Food, Agricultural and Environmental Sciences, Miyagi University, Miyagi, Japan; Present address: Laboratory of Food Biochemistry, Department of Bioscience, School of Agriculture, Tokai University, Kumamoto,Japan.

Chih‐Jung Kuo College of Veterinary Medicine, National Chung Hsing University, Taichung, Taiwan.

Jeffrey E. Lee Department of Laboratory Medicine and Pathobiology, Faculty of Medicine, University of Toronto, Toronto, Canada.

Alberto Marina Instituto de Biomedicina de Valencia, IBV‐CSIC, Valencia, Spain; CIBER de Enfermedades Raras (CIBERER‐ISCIII).

Lois E. Murray Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada.

Neil J. Oldfield Molecular Bacteriology and Immunology Group, School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, UK.

Christine Orengo Institute of Structural and Molecular Biology, University College London, Gower Street, London, UK.

Vijay Pancholi Department of Pathology, The Ohio State University College of Medicine and Wexner Medical Center, Columbus, Ohio, USA.

José R. Penadés Institute of Infection, Immunity and Inflammation, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK.

Melanie Pfeiffer Technical University Dresden, Dresden, Germany.

Christopher P. Ptak Department of Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA.

Nuria Quiles‐Puchalt Institute of Infection, Immunity and Inflammation, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK.

Manoj Raje Council of Scientific and Industrial Research (CSIR) Institute of Microbial Technology (CSIR‐IMTECH), Chandigarh, India.

Angela L. Riveroll Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada; Present address: Solarvest BioEnergy Inc. Summerville, PE, Canada.

Peter Robertson Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada; Present address: Canadian Food Inspection Agency, Dartmouth Laboratory, Dartmouth, Nova Scotia, Canada.

Natalia Salazar Laboratório de Bacteriologia, Instituto Butantan, São Paulo, Brazil.

Michael A. Sirover Department of Pharmacology, Temple University School of Medicine, Philadelphia, USA.

Fariza Shams Molecular Bacteriology and Immunology Group, School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, UK.

Patrick Trieu‐Cuot Unité de Biologie des Bactéries Pathogènes à Gram‐positif, Institut Pasteur, CNRS ERL 3526, 28 rue du Dr ROUX, Paris, France.

David P.J. Turner Molecular Bacteriology and Immunology Group, School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, UK.

Karla N. Valenzuela‐Valderas Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada.

Willem van Eden Department of Infectious Diseases and Immunology, Utrecht University, Utrecht, The Netherlands.

Karl G. Wooldridge Molecular Bacteriology and Immunology Group, School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, UK.

Masaya Yamaguchi Department of Oral and Molecular Microbiology, Osaka University Graduate School of Dentistry, Suita, Osaka, Japan.

Preface

Bacterial infection with exogenous pathogens can be thought of as a result of the evolution of specific bacterial behaviors that outwit the vast panoply of host immune defenses. Such interactions occur against the background of the vast colonization of Homo sapiens with a phylogenetically complex bacterial microbiota, whose interactions with the human host can only be guessed at. At its simplest, bacterial infection can be seen as a dynamic and evolutionarily constrained competition between the host and the genetically dynamic bacterial population of the environment. The major defining factor is bacterial virulence which could be defined simply as the population number required to infect a host organism. The fewer organisms required, the more virulent the organism. However, this has to be seen as a simplistic view of virulence, which is a systems‐based phenomenon with emergent properties. Virulence is a systems‐based concept which is dependent on the generation, by the bacterium, of molecules which can allow the bacterium to: (1) colonize; (2) survive the initial colonization process; (3) grow and, potentially, form biofilms; (4) defeat the approaches of the innate immune system; (5) deal with the adaptive immune cells; and finally (6) survive without killing the host.

The concept of virulence has given rise to the “virulence factor.” These will be best known in terms of the terrors of bacterial infection with gas gangrene (caused by Clostridium perfringen), the flesh‐eating bacterium (mainly describing Streptococcus pyogenes) with both pathologies being caused by enzymes, and flaccid and tetanic muscle spasms caused by Clostridium botulinum and Clostridium tetani toxins. Toxins are the main factor that comes to mind when thinking of bacterial virulence. However, they are only one of a range of molecules that aid the bacterium in its colonization and growth in the human organism. A range of other bacterial virulence factors include molecules which aid bacterial adhesion to matrices and cells, promote bacterial invasion of tissues and cells, control bacterial growth by binding essential metals, enable bacterial evasion of host immunity, and allow bacteria to enter low‐growth states (e.g., dormancy) which decrease their molecular signatures in the host. The formation of the bacterial biofilm involves a range of other virulence factors including those involved in quorum sensing, biofilm dispersion, and so on.

The renaissance of Bacteriology over the last 30 years (largely due to the upsurge in antibiotic resistance) has seen the identification of a wide range of bacterial virulence mechanisms and the discovery of a large number of molecularly distinct virulence factors, many of which are proteins. Since the early 1990s, it has become clear that among these distinct virulence proteins there exists a substantial number of proteins whose main function has nothing to do with bacterial virulence. Cytoplasmic proteins such as the glycolytic enzymes glyceraldehyde 3‐phosphate dehydrogenase (GAPDH) and enolase have been identified on the surface of a wide range of Gram‐positive and Gram‐negative bacteria, and have been reported to have a surprising number of diverse biological actions which are assumed to contribute to bacterial virulence. Indeed, where assessed using gene inactivation/upregulation, it has been established that these proteins have a direct role to play in bacterial virulence. These proteins are known as moonlighting proteins, which are defined as proteins with more than one unique biological action. Some of the bacterial moonlighting protein families from different bacterial species, although sharing >90% sequence identity, can produce quite distinct biological actions, thus increasing the virulence “range” of these proteins.

At the time of writing, around 90 bacterial proteins have been reported to exhibit more than one biological activity, with the moonlighting function being related to some virulence phenomenon. Many of these proteins are actually found in all three domains of life, and can therefore be thought of as shared signals. The discovery of the role of moonlighting proteins in bacterial interactions with their hosts reveals the plasticity of protein evolution as it relates to protein function and bacterial communication. Indeed, there are a small number of examples of human moonlighting proteins playing a role in enhancing bacterial colonization and virulence.

This book brings together the leading experts in the study of pathogenicity of bacterial moonlighting proteins. The book is divided into a number of related parts. In Part 1, the reader is introduced to the concept of protein moonlighting and is provided with current concepts in the evolution of protein moonlighting, its structural biological underpinnings, and its potential role in terms of cellular complexity and systems biology. Part 2 focuses on moonlighting in prokaryotes in a general sense, and includes chapters on general moonlighting proteins in bacterial infection and the role of moonlighting bacterial proteins in autoimmunity. Part 3 of this book discusses, in some detail, the various moonlighting proteins that are known to function as virulence factors, including: molecular chaperones and protein‐folding catalysts (Parts 3.1 and 3.2); glyceraldehyde 3‐phosphate dehydrogenase (GAPDH) (Part 3.3); enolase (Part 3.4); other glycolytic enzymes (Part 3.5); other metabolic enzymes (Part 3.6); miscellaneous proteins (Part 3.7); and bacterial moonlighting proteins that function to bind cytokines (Part 3.8). Finally, in Part 3.9 the subject switches to novel findings in bacteriophage and virus biology in which moonlighting proteins are able either to promote bacterial virulence or aid in viral infection.

This book will be of interest to a range of scientists. Clearly the major reader will be the bacteriologist/cell biologist interested in the mechanisms of bacterial virulence and in the possible role of bacterial moonlighting proteins as therapeutic targets. Given the widespread use of some of these moonlighting proteins as virulence determinants by many of the bacterial pathogens of Homo sapiens, this is a possibility. Other readers will include immunologists, biochemists, molecular biologists, and pathologists focusing on the biology of the cell stress response and those interested in the diversity of protein structure and function. The finding that bacteria encode high‐affinity binding proteins for key proinflammatory cytokines such as IL‐1β and TNFα also suggests a therapeutic potential for such molecules.

About the Editor

Brian Henderson is Professor of Biochemistry in the Department of Microbial Diseases at the UCL-Eastman Dental Institute, University College London. He has worked in academia, both in the UK and North America, and also in the pharmaceutical and biopharmaceutical industry. He has been a cell biologist, immunologist, and pharmacologist and over the past 20 years has focused on bacteria–host interactions in relation to human infection and the maintenance of the human microbiota. This is known as the discipline of Cellular Microbiology, and Henderson published the first book on this subject in 1999. At the inception of his career as a cellular microbiologist, he discovered a potent bone-destroying protein secreted by a pathogenic oral bacterium. This protein turned out to be the cell stress protein, heat-shock protein (Hsp)60. This was one of the earliest bacterial moonlighting proteins discovered, and is the reason that the editor has spent the last 20 years exploring the role of protein moonlighting in the life of the bacterium and its interactions with its human host. Henderson has written or edited 17 books and monographs and was the senior editor of the Cambridge University Press monograph series Advances in Molecular and Cellular Microbiology. His last book, published in 2013, was entitled Moonlighting Cell Stress Proteins in Microbial Infections.

Part IOverview of Protein Moonlighting

1What is Protein Moonlighting and Why is it Important?

Constance J. Jeffery

Department of Biological Sciences, University of Illinois at Chicago, Chicago, Illinois, USA

1.1 What is Protein Moonlighting?

Moonlighting proteins exhibit more than one physiologically relevant biochemical or biophysical function within one polypeptide chain (Jeffery 1999). In this class of multifunctional proteins, the multiple functions are not due to gene fusions, multiple RNA splice variants or multiple proteolytic fragments. The moonlighting proteins do not include pleiotropic proteins, where a protein has multiple downstream cellular roles in different pathways or physiological processes that result from a single biochemical or biophysical function of a protein. Moonlighting proteins also do not include families of homologous proteins if the different functions are performed by different members of the protein family.

Some of the first moonlighting proteins to be identified were taxon‐specific crystallins in the lens of the eye. These proteins, including the delta 2 crystallin/arginosuccinate lyase in the duck (Wistow and Piatigorsky 1987), upsilon crystallin/lactate dehydrogenase A in the duckbill platypus (van Rheede et al. 2003), eta‐crystallin/cytosolic aldehyde dehydrogenase (ALDH class 1) in the elephant shrew (Bateman et al. 2003), and several others, are ubiquitous soluble enzymes that were adopted as structural proteins in the lens. Other well‐known moonlighting proteins include soluble enzymes in biochemical pathways that also bind to DNA or RNA to regulate transcription or translation. Human thymidylate synthase (TS), a cytosolic enzyme in the de novo synthesis of the DNA precursor thymidylate, also binds to mRNA encoding TS to inhibit translation (Chu et al. 1991). The Salmonella typhimurium PutA protein is an enzyme with proline dehydrogenase and proline oxidase pyrroline‐5‐carboxylic acid dehydrogenase activity when it is bound to the inner side of the plasma membrane (Menzel and Roth 1981a, b), but it also binds to DNA and moonlights as a transcriptional repressor of the put operon (Ostrovsky de Spicer et al. 1991; Ostrovsky de Spicer and Maloy 1993). The E. coli BirA biotin synthase is an enzyme in the biotin biosynthetic pathway that is also a bio operon suppressor (Barker and Campbell 1981). Saccharomyces cerevisiae N‐acetylglutamate kinase/N‐acetylglutamyl‐phosphate reductase (Arg5,6p) is an enzyme in the arginine biosynthetic pathway (Boonchird et al. 1991; Abadjieva et al. 2001) and also binds to mitochondrial and nuclear DNA to regulate expression of several genes (Hall et al. 2004). Kluyveromyces lactis galactokinase (GAL1) phosphorylates galactose and is also a transcriptional activator of genes in the GAL operon (Meyer et al. 1991).

Perhaps even more surprising than the fact that some proteins can perform such different functions is that such a large variety of proteins moonlight. Over the past few decades, hundreds of proteins have been shown to moonlight (Mani et al. 2015; moonlightingproteins.org). They include many types of proteins: enzymes, scaffolds, receptors, adhesins, channels, transcription and translation regulators, extracellular matrix proteins, growth factors, and many others. They are active in a variety of physiological processes and biochemical pathways, are found in the cytoplasm, nucleus, mitochondria, on cell surface, and other cellular compartments, and some are secreted. They are also expressed in many different cell types within a species. They are found in a variety of species from throughout the evolutionary tree. They are common in eukaryotes in humans and other placental and monotreme (i.e., platypus) mammals, reptiles, birds, amphibians, fish, worms, insects, plants, fungi, and protozoans. A few are found in archea and many more have been identified in eubacteria, including pathogenic species (Clostridium difficile, Helicobacter pylori, Pseudomonas aeruginosa, Staphylococcus, etc.) as well as nonpathogenic, commensal bacteria, including health‐promoting or “pro‐biotic” species (Bifidobacterium). A few moonlighting proteins have even been found in viruses.

The variety also extends to the combinations of functions that are observed. Many of the known moonlighting proteins are cytosolic enzymes, chaperones, or other proteins that exhibit a second function in other cellular locations, for example as a receptor on the cell surface. Several proteins described in more detail in other chapters are cytosolic enzymes or chaperones that are secreted to serve as growth hormones or cytokines. For example, an enzymatic function and an extracellular cytokine function are found in phosphoglucose isomerase/autocrine motility factor (Gurney et al. 1986a, b; Chaput et al. 1988; Faik et al. 1988; Watanabe et al. 1996; Xu et al