JB Accepts, published online ahead of print on 30 November 2012 J. Bacteriol. doi:10.1128/JB.01743-12 Copyright © 2012, American Society for Microbiology. All Rights Reserved.

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ClpL is required for folding of CtsR in Streptococcus mutans

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Liang Tao and Indranil Biswas* Department of Microbiology, Molecular Genetics and Immunology, University of Kansas Medical Center, 3901 Rainbow Boulevard, Kansas City, KS 66160 Running Title: ClpL acts as a chaperone *Corresponding author. Mailing address: 2032 BERI, 3901 Rainbow Blvd. MSC 3029. Kansas City, KS 66160. Phone: (913) 588-7019 Fax: (913) 588-7295 E-mail: [email protected]

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SUMMARY

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ClpL, a member of HSP100 family, is widely distributed in gram-positive bacteria but is

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absent in gram-negative bacteria. Although ClpL is involved in various cellular

20

processes such as stress tolerance response, long-term survival, virulence, and

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antibiotic resistance; the detailed molecular mechanisms are largely unclear. Here

22

we report that ClpL acts as a chaperone to properly fold CtsR, a stress-response

23

repressor, and prevents it from forming protein aggregates in Streptococcus mutans.

24

In vitro, ClpL was able to successfully refold urea-denatured CtsR but not aggregated

25

proteins. We suggest that ClpL primarily recognize soluble but denatured substrates

26

and prevent formation of large protein aggregates.

27

C-terminal D2-small domain of ClpL is essential for the observed chaperone activity.

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Since ClpL widely contributes to various cellular functions, we speculate that ClpL

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chaperone activity is necessary to maintain the cellular homeostasis.

We also found that in vivo

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Introduction

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Bacteria are constantly exposed to stressful environments such as exposure to free

33

radicals, acidic or alkaline conditions, or high temperature. Exposure to environmental

34

stresses often causes denaturation of cellular proteins that subsequently accumulate

35

within the cell as protein aggregates. To encounter the detrimental effect of thermal

36

and other stresses, bacteria transiently synthesize a highly conserved set of proteins

37

with molecular chaperone or protease activities and are generally referred to as heat

38

shock proteins (HSPs) [1,2]. HSPs are ubiquitous in bacteria and depending on the

39

molecular weight HSPs are grouped into four major classes: HSP100, HSP70, HSP60,

40

and small HSP families [1,3]. HSP100 subgroup, which is also known as caseinolytic

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protease system (Clp), is typically an AAA+ (ATPases associated with a variety of

42

cellular activities) super-family protein that often forms complexes with a peptidase

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subunit, such as ClpP, for proteolytic activity required for removing damaged and

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denatured proteins, as well as protein folding functions [4,5]. Clp proteases also play

45

important roles in regulating various cellular functions such as controlling growth at low

46

or high temperature, competence development, sporulation, and virulence [6,7,8,9].

47

According to the number of ATP-binding domains on the polypeptide chain,

48

regulatory ATPases subunits can be grouped into class-I (two ATP-binding domains,

49

AAA-1 and AAA-2) and class-II (one ATP-binding domain, AAA-1) [10]. Clp proteins

50

have also been categorized into various classes based on the length of the spacer

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sequence at the middle region, overall sequence similarity, and variation in the N- and

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C-terminal regions [10]. Clp ATPases that interact with ClpP peptidase, such as ClpA

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and ClpX, encode a conserved IGF motif that is present at a surface loop [11,12].

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These ATPase are assembled into hexameric rings with a narrow pore in the center

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and use energy generated by the ATP hydrolysis to unfold target protein substrates

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and translocate them into the chamber of associated barrel-like ClpP protease

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complex, where the peptide bonds are cleaved [13,14,15,16]. There are a few Clp ATPases that do not interact with ClpP peptidase; instead they

59

sometimes cooperate with the HSP70 system to function as a chaperone that can

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disaggregate and refold denatured proteins [17]. In Escherichia coli, ClpB is one such

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Clp ATPase that does not interact with ClpP. ClpB contains two ATP-binding domains

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(AAA-1 and AAA-2) that are separated by a middle domain (M-domain) forming a

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coiled-coil structure [18,19]. Like other Clp ATPases, ClpB also forms a hexameric-ring

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structure and cooperates with DnaK chaperone system that includes DnaK, DnaJ, and

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GrpE [17,19]. ClpB has a remarkable ability to rescue proteins from an aggregated

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state by aiding the disaggregation of denatured proteins. The complete refolding of the

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denatured proteins is dependent on the concerted effort of the cognate

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DnaK-DnaJ-GrpE system [20,21,22]. ClpB alone may also suppress the aggregation

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of labile proteins with its monomeric or dimeric form [23]. Despite a wealth of

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biochemical and structural information, the mechanistic aspects of this bi-chaperone

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system remain poorly understood. This is in part because ClpB and DnaK system

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interact only transiently [24,25] and recognize substrate proteins via sequential binding

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[26]. Recently, it was suggested that ClpB M-domain determines the specificity

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between ClpB and DnaK cooperation, which is required for protein disaggregation and

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thermo tolerance [27,28]. In many low-GC gram-positive bacteria, in addition to ClpB, another protein, ClpL,

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with homology to ClpB is also present [6,29]. In Streptococcus pneumoniae, a

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respiratory pathogen, ClpL has been shown to be involved in thermo tolerance, acid

79

tolerance, and virulence regulation, and resistance to various antibiotics that target the

80

cell-wall [29,30,31,32].

81

Streptococcus mutans, a primary etiological agent of dental caries, possesses five

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proteins that belong to the Clp family including ClpB, ClpC, ClpE, ClpL, and ClpX [33].

83

CtsR, a major repressor for the Clp ATPases [34,35], mainly regulates the expression

84

of stress response genes by recognizing a tandemly repeated hepta-nucleotidic

85

sequence known as CtsR-box [35]. We recently demonstrated that CtsR, is

86

accumulated in higher amounts in cells that do not have a functional ClpL [36]. The

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finding was interesting since in most gram-positive bacteria CtsR is degraded by

88

ClpCP [37]. In this study, we showed that in the absence of ClpL, CtsR protein is

89

accumulated largely in the S. mutans cells as inactive aggregated form. Direct

90

protein-protein interaction between ClpL and CtsR was confirmed both in vitro and in

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vivo. We also observed that the presence of ClpL largely enhanced the refolding of

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urea-denatured CtsR in vitro. In vivo complementation showed that ClpL lacking the

93

D2-Small domain was no longer functional to alleviate the CtsR aggregates. These

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results indicate that ClpL is crucial for proper folding of CtsR protein in S. mutans even

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at ambient growth temperature.

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MATERIALS AND METHODS

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Bacterial strains, plasmids, and growth conditions

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The S. mutans strains and plasmids used in this study are listed in Table 1.

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Escherichia coli strains were routinely grown in Luria-Bertani medium supplemented

101

with (when necessary) 100 μg/ml ampicillin, and/or 50μg/ml kanamycin. S. mutans

102

isolates were normally grown at 37°C in Todd-Hewitt medium (BBL, BD)

103

supplemented with 0.2% yeast extract (THY medium). When necessary, 5 μg/ml

104

erythromycin and/or 400 μg/ml kanamycin was included in THY medium.

105 106

Protein extraction and western blot analysis

107

For S. mutans protein extraction, unless otherwise stated, overnight grown cultures

108

were inoculated in THY medium and grown to exponential phase (optical density at

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600 nm [OD600] = 0.4). A 10 ml aliquot was harvested by centrifugation, resuspended

110

in 600 μl of phosphate-buffered saline (PBS), and homogenized with a bead-beater

111

(MP Biomedicals, LLC). Cell lysate was centrifuged at 18000 g for 10 minutes and

112

~200 μl of supernatant was carefully transferred into a new tube and kept as the

113

soluble fraction. The remaining supernatant was removed and the cell debris (pellet)

114

together with silicon beads were washed three times with PBS and resuspended in

115

200 μl of PBS and stored as the insoluble fraction. Both soluble and insoluble

116

fractions were added with protein sample buffer, boiled for 5 minutes, and separated

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by SDS-PAGE. The gels were stained by Coomassie blue R-250 or blotted onto

118

PVDF membranes. Western blot assays were carried out using standard techniques.

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An anti-polyhistidine (anti-His) monoclonal antibody (Sigma) was used as primary

120

antibody to detect his-tagged proteins. For western blot experiments using the cell

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lysate as samples, the abundance of cellular enolase was chosen as an internal

122

control. PVDF membranes were stripped with Tris-buffered saline (TBS) containing

123

2% SDS and 100 mM β-mercaptoethanol, and reprobed by an anti-S. mutans enolase

124

(anti-SmuEno) polyclonal (Genscript) antibody. Western blots were developed with

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Pierce ECL plus reagents (Thermo Scientific); and the fluorescent signals were

126

detected by Typhoon FLA9000 biomolecular imager (GE Healthcare).

127

blot experiments were repeated as least twice to confirm the results.

All western

128 129

Construction of PclpP-gusA reporter strain and β-glucuronidase (Gus) assay

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Plasmid pIB521 containing PclpP-gusA fusion was previously used to measure clpP

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promoter activity [38]. Plasmid pIB521 was linearized with BglI, transferred to S.

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mutans IBSJ3 via natural transformation, using a protocol described previously [39] to

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generate strain IBSJ9.

134

Briefly, the optical density at 600 nm (OD600) of exponential phase culture was

135

recorded before cell harvesting. One ml of culture was harvested, washed in saline,

136

and resuspended in 500 μl of Z-buffer (60 mM Na2HPO4, 40 mM Na2HPO4, 10 mM

137

KCl, 1 mM MgSO4, and 20 mM dithiothreitol [DTT]). Cells were homogenized by bead

138

beating. 200μl of cell lysate was then transferred to a new tube, 100μl of

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p-nitrophenyl-β-D-glucoside (4 mg/ml in Z-buffer) was added, and incubated in 37°C

140

until a yellow color developed. The reaction was stopped by addition of 200 μl of 1 M

Gus assay was performed as previously described [36].

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Na2CO3. The absorbance at 420 nm (A420) and the time period of the reaction in

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minute (T) were noted. Gus activity was defined as [1000×A420]/[T ×OD600] in Miller

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units (MU).

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Co-expression of ClpL and CtsR in E. coli

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The open reading frame (ORF) of ctsR was amplified from UA159 genomic with

147

primers

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ctsR-duet-R (5’-CTTCTCGAGTCATAGATGGTATCCTTTTCTATC-3’); and restricted

149

with NdeI-XhoI. Similarly, the ORF of clpL was amplified from UA159 genomic with

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primers clpL-duet-F (5’-CTTGGATCCGATGGCAAATTTTAATGGACGCG-3’) and

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clpL-duet-R

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restricted with BamHI-PstI. The restricted ctsR and clpL fragments were cloned into

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vector pETduet, respectively, to create pIBJ33 and pIBJ34. The restricted ctsR

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fragment was then cloned into NdeI-XhoI digested pIBJ34 to obtain pIBJ35. Plasmid

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pIBJ35 was transformed into E. coli BL21 (DE3) and allowed to co-express two

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proteins under the inducement of Isopropyl-β-D-thio-galactoside (IPTG).

ctsR-duet-F

(5’-CTTCATATGACGTCAAAAAATACTTCAG-3’)

(5’-CTTCTGCAGTTAAGCTTCTTCAATAATCAATTTGTC-3’),

and

and

157 158

Protein purification and His-tag pull-down assay

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His-CtsR expression was induced with 200 μg/L anhydrotetracycline in E. coli DH5α

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with pIBC37. E. coli BL21 (DE3) with either pIBJ33 or pIBJ34 was used for the

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over-expression of His-ClpL or CtsR-Stag induced by 1 mM IPTG. His-tagged proteins

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were purified by nickel-nitrilotriacetic acid (Ni-NTA) resin (Novagen) in accordance

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with the manufacturer’s instructions. The protein was dialyzed overnight against a

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buffer containing 20 mM Tris-Cl (pH 7.4), 100 mM NaCl and 1 mM DTT. Soluble

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CtsR-Stag protein was extracted from purified inclusion bodies with PBS containing 8

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M Urea. The protein extract was then dialyzed against PBS with Urea of gradient

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concentrations and final protein solution contained 2 M urea. The purity of the

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proteins was >95%, as determined by SDS-PAGE analysis. Protein concentrations

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were

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Spectrophotometer (Thermo Scientific).

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Pull-down assay for His-tagged proteins was performed using standard method.

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Briefly, 50 μg of His-ClpL and/or 5 μg of CtsR-Stag were added to 1 ml of binding

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buffer (20 mM sodium phosphate, 150 mM NaCl, 4 mM ATP; pH 7.6) together with 30

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μl of pre-washed settled Ni-NTA resin. The mixture was incubated at 4 °C with rotation

175

for 30 minutes; resin was washed twice with PBS containing 30 mM of imidazole

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followed by washing twice with PBS containing 0.01% Triton-X 100.

177

were eluted with elution buffer (50 mM sodium phosphate, 300 mM NaCl, 250 mM

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imidazole; pH 7.4) and subjected to SDS-PAGE analysis.

estimated

by

the

absorbance

at

280

nm

using

Nanodrop

2000c

Bound proteins

179 180

Renaturation of S. mutans CtsR

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CtsR-Stag dissolved in PBS containing 2 M urea was used for refolding assay by

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dialysis at 4°C.

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of CtsR-Stag before dialysis. To initiate renaturation, 400 μg/ml CtsR-Stag was

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dialyzed at 4°C against a dialysis buffer containing 20 mM Tris-Cl (pH 7.4), 50 mM

Urea (2 M) was pre-added to all additives to prevent the precipitation

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NaCl, 1 mM DTT, 10% glycerol, 4 mM ATP in the presence or absence of His-ClpL of

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varying concentrations. The dialyzed samples were centrifuged at 18000 g for 10 min,

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and the supernatant fractions were analyzed by SDS-PAGE. The efficiency of

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renaturation was determined by measuring the intensity of protein bands by

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ImageQuantTL software on Coomassie blue stained gel. The EMSA assay used for

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verifying the DNA binding activity of the refolded CtsR-Stag was performed as

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previously described [36]. To test the potential for folding CtsR aggregates by ClpL,

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insoluble CtsR-Stag aggregates were incubated with His-ClpL in the refolding buffer

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(20 mM Tris-Cl, 50 mM NaCl, 1 mM DTT, 10% glycerol, 4 mM ATP; pH 7.4) for 4

194

hours at 37 °C. The mixture was then centrifuged at 18000 g for 10 minutes and the

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supernatant fraction was separated by SDS-PAGE and transferred to a PVDF

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membrane. The detection of CtsR-Stag was performed by western blot using a

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monoclonal anti-Stag antibody (Novagen).

198 199

Complementation of clpLΔN and clpLΔC

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A DNA fragment encoding clpL ORF but lacking the 117 N-terminal amino residues

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was amplified from UA159 genomic DNA using primers ClpLΔN-184km-F

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(5’-CACGGATCCATGCCTGTTCTGGTCGGTGATG-3’)

203

(5’-CACGGTACCACAGCTTCTTCAATAATCAATTTGTC-3’).

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fragment encoding clpL ORF but lacking the 78 C-terminal amino residues was also

205

amplified

206

(5’-CACGGATCCATGGCAAATTTTAATGGACGC-3’)

from

UA159

genomic

DNA

using

and

Similarly,

primers and

ClpL-184km-R a

DNA

ClpL-184km-F ClpLΔC-184km-R

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(5’-CACGGTACCACGTGAGAGAATTCAATAACTGC-3’). The amplified fragments

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were cloned into vector pIB184Km [40], which contains the P23 promoter, to create

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pIBJ63 and pIBJ64, respectively. Strain IBSJ3/pIBJ1 was transformed with either

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pIBJ63 or pIBJ64, and the transformants were selected on THY agar plates

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containing both erythromycin and kanamycin. The presence of the complementary

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genes and his-ctsR was confirmed on the selected transformants by PCR.

213 214

Bacterial Adenylate Cyclase Two-Hybrid (BACTH) assay

215

Two compatible plasmids, one expressing the T18 fusion (PUT18, Euromedex) and

216

the other expressing the T25 fusion (PKNT25, Euromedex), were chosen for our

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BACTH assays. DNA fragments encoding full-length clpL, clpL lacking 117 N-terminal

218

amino acids (ClpLΔN), or clpL lacking 78 C-terminal amino acids (ClpLΔN) were

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amplified from the UA159 genomic DNA and cloned into vector pUT18 to create

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pIBJ58, pIBJ59 and pIBJ60, respectively. Meanwhile, the DNA fragments encoding

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ctsR or clpL were amplified and cloned into vector pKNT25 to create pIBJ55 and

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pIBJ62, respectively. Primers used for above gene amplifications are as follows:

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ClpL-T18-F

224

(5’-CACGGTACCACAGCTTCTTCAATAATCAATTTGTC-3’),

ClpLΔN-T18-F

225

(5’-CACGGATCCACCTGTTCTGGTCGGTGATG-3’),

ClpLΔC-T18-R

226

(5’-CACGGTACCACGTGAGAGAATTCAATAACTGC-3’),

227

(5’-CACGGATCCAATGACGTCAAAAAATACTTCAG-3’),

228

(5’-CACGGTACCACTAGATGGTATCCTTTTCTATC-3’).

(5’-CACGGATCCAATGGCAAATTTTAATGGACGC-3’),

ClpL-T18-R

CtsR-T25-F and

CtsR-T25-R

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Two compatible plasmids, a pUT18 derivative and a pKNT25 derivative, were

230

co-transformed into E. coli indicator strain BTH101 by electroporation and screened

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on LB agar plates containing both ampicillin and kanamycin. The presence of the

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target genes in the selected transformants was verified by PCR. The confirmed

233

bacterial cells were streaked onto the LB agar plates containing antibiotics, IPTG (0.5

234

mM), and 5-bromo-4-chloro-3-indolyl-β-D- galactopyranoside (X-Gal, 40 μg/ml) and

235

grown at 30 °C. If there is an interaction between the two proteins of interest, the

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colonies turn blue within 24-72 hours according to the manufacturer’s instruction.

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Quantification of the functional complementation mediated by the interaction of two

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proteins of interest was achieved by measuring β-galactosidase activity.

239 240

β-galactosidase assay

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E. coli BTH101 cells with plasmids of interest were inoculated into liquid LB medium

242

containing ampicillin, kanamycin and IPTG (0.5 mM). The culture was grown

243

overnight at 37 °C to reach the stationary phase. The OD600 of the overnight culture

244

was recorded before harvesting. One milliliter of overnight culture was centrifuged;

245

cell pellets were washed twice with PBS and resuspended in the equal volume of

246

Z-buffer. 100 μl of resuspended bacterial cells were diluted in 1 ml of Z-buffer (dilution

247

factor [DF] =10). Afterwards, 100 μl of chloroform and 50 μl of 0.1% SDS were added

248

and mixed well to permeabilize the cells. 250 μl of mixture was then transferred to a

249

new

250

ο-nitrophenyl-β-galactoside (4 mg/ml in Z-buffer) was added, and incubated at 28°C

microfuge

tube

and

brought

to

28°C,

50

μl

of

pre-warmed

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229

251

until a yellow color developed. The reaction was stopped by the addition of 200 μl of 1

252

M Na2CO3. The A420 and the precise time period of the reaction in minute (T) were

253

recorded. β-galactosidase activity was defined as [1000×A420×DF]/[T×OD600] in Miller

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units (MU).

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256

RESULTS

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CtsR is accumulated in ΔclpL S. mutans strain as inactive aggregated form

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Our previous study showed that CtsR protein was accumulated in large amount in a

259

clpL-deficient S. mutans strain [36].

260

accumulated CtsR (His-CtsR) is predominantly present in the pellet fraction and not in

261

the soluble fraction of the cell lysate from the ΔclpL strain (IBSJ3/pIBJ1). On the other

262

hand, very little His-CtsR protein was found in the insoluble fraction in the wild-type

263

background (i.e. UA159/pIBJ1; Fig. 1A).

264

specific to CtsR, we used HcrA, a transcriptional repressor also involved with heat

265

shock response, as control.

266

wild type and the mutant was similar, suggesting that ClpL is not involved with the

267

protein accumulation.

268

mutans cells and most of the protein was present as aggregated form, suggesting that

269

the accumulated form may be functionally inactive.

270

To determine whether the accumulation of CtsR in ΔclpL cells correlates with CtsR’s

271

ability to repress transcription, we used a reporter fusion strain that carry PclpP-gusA

272

construct in the chromosome.

273

binding and PclpP-gusA was successfully used to measure the CtsR repressor

274

activity [36,38]. Since CtsR is a repressor, low amounts of active CtsR would produce

275

increased Gus activity from this promoter fusion. PclpP-gusA fusion construct was

276

introduced into the wild type S. mutans strain UA159 and the ΔclpL mutant strain

277

IBSJ3 to create strains IBS514 and IBSJ9, respectively, and β-glucuronidase (GusA)

Further analyses suggested that the

As shown in Fig. 1B, the amount of HrcA protein in the

In contrast, CtsR was greatly accumulated in the ΔclpL S.

The clpP promoter is an authentic target for CtsR

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To determine if the accumulation was

278

activity was measured from these reporter strains (Fig. 1C). Surprisingly, we observed

279

that the GusA activity was increased in IBSJ9 as compare with IBS514 even though

280

the amount of CtsR protein was much higher in the ΔclpL mutant strain (Fig. 1A).

281

These results suggest that although the CtsR protein has been accumulated in the

282

cell, in the absence of ClpL the accumulated CtsR remains as inactive form.

284

ClpL does not directly degrade CtsR in vitro

285

While aggregated proteins are usually highly refractory to various cellular proteases,

286

we wanted to investigate whether ClpL directly participates in the degradation of CtsR

287

protein or ClpL prevents misfolding of CtsR and thereby reducing the total

288

aggregation in the cells. Sequence analysis suggested that ClpL does not harbor any

289

peptidase-like domains and unlike other Clp ATPase proteins (such as ClpC or ClpX),

290

clpL does not encode IGF motif that is required for interaction with ClpP [11].

291

Therefore, it is unlikely that ClpL directly involved in the degradation of native CtsR to

292

control the protein level.

293

proteolytic potential of ClpL. Purified His-CtsR (100 ng/ml) was incubated with

294

His-ClpL (300 ng/ml) and whole cell lysate (500 μg/ml) from S. mutans UA159 in the

295

presence of 4 mM ATP at 37°C. The mixture was then separated by sodium dodecyl

296

sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a

297

polyvinylidene difluoride (PVDF) membrane. Both His-CtsR and His-ClpL were

298

detected with anti-His antibody. No obvious degradation of His-CtsR was observed

299

after 30 minutes incubation (data not shown). Therefore, we speculate that ClpL

We designed an in vitro degradation assay to test the

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283

300

controls the CtsR level in S. mutans by preventing the protein aggregation formation

301

rather than degradation of the natural form CtsR protein.

302

ClpL interacts with CtsR both in vitro and in vivo

304

If ClpL were to engage in the folding of CtsR protein, one would expect ClpL to

305

interact directly with CtsR.

306

and CtsR, we employed an in vitro pull-down assay with purified proteins to assess

307

the affinity between these two proteins.

308

C-terminal tagged CtsR (CtsR-Stag) were expressed separately and purified from E.

309

coli as bait and prey proteins.

310

exhibited a strong binding affinity to His-ClpL while it had no binding to empty Ni-NTA

311

resin (Fig. 2A).

312

The ClpL-CtsR interaction was also verified in vivo.

313

His-ClpL alone in E. coli cells. As expected, majority of the ClpL was in the soluble

314

fraction when induced at 37°C. On the other hand when CtsR-Stag alone was

315

expressed E.coli cells, the protein was always in the insoluble fraction even when the

316

cells were grown at a lower temperature. However, when both His-ClpL and

317

CtsR-Stag were co-expressed in E. coli, we found that nearly all His-ClpL protein

318

became insoluble (Fig. 2B). This data indicate that the ClpL-CtsR interaction also

319

exist in vivo. To further confirm the in vivo protein-protein interaction between ClpL

320

and CtsR, a bacterial two-hybrid (BACTH, Euromedex) system was employed to test

321

their binding affinity.

To explore the possibility of the interaction between ClpL

N-terminal tagged ClpL (His-ClpL) and

CtsR-Stag (dissolved in PBS containing 2 M urea)

For this, we first expressed

Bacterial colonies that co-expressed ClpL-T18 and CtsR-T25

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303

322

fusion proteins became light blue, while the control bacterial colonies remained white

323

(Fig. 2C).

324

and is shown in Fig. 2D. These data suggest that ClpL interacts directly with CtsR,

325

however the interaction between fusion proteins appears to be not very strong since

326

the β-gal activity was low to moderate.

The interaction affinity was quantified using β-galactosidase (β-gal) assay

328

ClpL helps folding of CtsR in vitro

329

Since we determined the protein-protein interaction between ClpL and CtsR, we

330

speculated that ClpL might have a chaperone activity that helps folding of CtsR.

331

Note that, CtsR was hardly soluble when expressed in E. coli cells despite the

332

addition of an N-terminal or a C-terminal tag; and was mainly inactive when

333

accumulated in ΔclpL S. mutans strain. Urea was used to solubilize CtsR-Stag

334

aggregates and we examined the chaperone activity of ClpL. When CtsR-Stag was

335

dialyzed alone, soluble CtsR protein was hardly detected after urea removal (Fig. 3).

336

Addition of sheared salmon sperm DNA, which previously reported to help refolding of

337

HrcA [41], had little contribution on CtsR-Stag refolding (data not shown). However,

338

when twice the molar amount of His-ClpL was supplemented during the dialysis, the

339

refolding efficiency of CtsR-Stag was attained to approximately 80% (Fig. 3B).

340

refolded CtsR protein, which was devoid of urea, was active and retained its target

341

DNA binding activity (Fig. 3C).

342

the aggregate formation or can disaggregate once the protein aggregates are formed,

343

we incubated His-ClpL with CtsR-Stag aggregates in the presence of ATP and without

This

To test whether ClpL helps the folding of CtsR before

17

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327

344

urea.

No soluble CtsR-Stag was detected in the supernatant fraction after 4 hours of

345

incubation, suggesting that ClpL alone was not sufficient to resolubilize the CtsR

346

aggregates (Fig. 3D).

347

well as in vivo and prevent accumulation of CtsR aggregates due to proper folding the

348

protein in the cell.

However, our data suggest that ClpL can fold CtsR in vitro as

350

Both N- and C- terminal domains contribute to oligomerization of ClpL

351

Primary sequence analysis showed that S. mutans ClpL contained two highly

352

conserved ATP-binding regions (AAA-1 and AAA-2 domain) and a D2-small domain at

353

its C-terminal domain. By comparison protein sequences of known Clp ATPases in S.

354

mutans, we found that the domain organization of ClpL is similar to ClpB, which is

355

widely present in both gram-positive and gram-negative bacteria (Fig. 4A). ClpB is

356

self-assembled to form an oligomeric complex; mainly as hexamer but also exist as

357

monomer and dimer [18,23]. When we applied the purified ClpL protein onto a

358

Superdex 200 10/300 GL size exclusion column, the elution profile yielded two equal

359

intensity peaks that correspond to molecular weights of >600 KDa and ~470 KDa

360

(data not shown). The latter size corresponds to a hexameric protein and the larger

361

size

362

(tris[2-carboxyethyl]phosphine, a reducing agent), greatly reduced the intensity of the

363

larger peak, but did not abolish it (data not shown).

364

ClpB also exists as hexamer, and perhaps other higher order oligomers.

365

the sequence similarity with ClpB and the presence of the C-terminal D2-small

indicates

protein

aggregates.

Treating

the

sample

with

TCEP

Thus, it appears that ClpL, like Because of

18

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349

366

domain that forms a tight interface with the AAA2 domain of the neighboring subunit

367

of ClpB [18], we

368

domains of ClpL in oligomerization. Towards this end, truncated ClpL proteins with

369

either N- or C-terminal deleted regions were evaluated in BATCH assay for

370

interactions (Fig. 4B).

371

interacted strongly with ClpL-T25 fusion (Fig. 4C). We also observed that C-terminal

372

D2-small domain is essential for ClpL oligomerization, since deleting this domain in

373

ClpL-T18 (ClpLΔC-T18) resulted in the loss of interaction with ClpL-T25 (Fig. 4C).

374

However, deleting N-terminal 117 residues in ClpL-T18 (ClpLΔN-T18) also impaired

375

its interaction with ClpL-T25 (Fig. 4C). This is in contrast to ClpB protein where

376

deletion of N-terminal domain does not abolish the oligomerization as reported

377

previously [42]. Taken together, our data indicate that ClpL can form hexamer, and

378

both N- and C-terminal domains contribute to the protein oligomerization.

379

observed that deletion of N- or C-terminal domains of ClpL also weakens the

380

interaction with CtsR (Fig. 2C and D).

then wanted to explore the contribution of N- and C-terminal

As expected, the assay showed that ClpL-T18 fusion

381 382

C-terminal domain of ClpL is important for chaperone activity

383

To further understand the contribution of ClpL N-terminal and C-terminal domains to

384

its chaperone activity. We expressed either ClpLΔN or ClpLΔC in our ΔclpL mutant

385

strain IBSJ3/pIBJ1. Our previous study showed that His-CtsR accumulation was

386

prevented in IBSJ3/pIBJ1 when the full length ClpL was expressed in trans from a

387

plasmid [36].

We found that the expression of ClpLΔN in IBSJ3/pIBJ1 can also

19

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We also

388

prevent the cellular accumulation of His-CtsR (Fig. 5). However, level of His-CtsR

389

accumulation was unchanged when ClpLΔC was expressed in IBSJ3/pIBJ1 (Fig. 5),

390

suggesting that deletion of C-terminal D2-small domain in ClpL resulted in the loss of

391

its chaperone activity. Thus, C-terminal D2 domain appears to play an important role

392

in protein folding.

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393 394

20

DISCUSSION

396

ClpL is a unique member of the HSP100 family that does not encode the motif

397

required for interaction with ClpP. This protein is strictly found in gram-positive

398

bacteria including streptococci and is involved in various cellular processes such as

399

stress tolerance response, virulence, long-term cell survival, and antibiotic resistance

400

[7,30,31,32].

401

functions of this protein.

402

regulator, is accumulated in the clpL-deficient but not in the clpP-deficient strain [36].

403

Subsequently, we found that most of the accumulated protein was present in the

404

pellet fraction.

405

increased in the clpL-deficient strain, the accumulated CtsR protein seemed to be

406

improperly folded in the mutant strain.

407

pneumoniae ClpL can refold in the presence of ATP urea-denatured rhodanese (a

408

non-streptococcal protein) in a dose-dependent manner [29], indicating that ClpL

409

might have a chaperone activity.

410

involved in folding of CtsR in the cell.

411

showed that CtsR is a bona fide substrate for ClpL.

Despite its importance, very little is known about the molecular Our earlier study indicated that CtsR, a major heat shock

Since the expression from a CtsR-repressed promoter (PclpP) was

A previous study reported that S.

Therefore, we hypothesized that ClpL might be Our study presented here conclusively

412 413

In the wild type S. mutans cells, CtsR level is very low even after over expression of

414

CtsR from a multicopy plasmid indicating that the protein is readily degraded in the

415

cell. Our data showed that ClpL did not enhance the degradation of CtsR protein in its

416

native form; thus we suggest that the degradation step is correlated with protein

21

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395

417

folding stage. This step may require involvement of other accessory proteins.

Since

418

ClpL does not contain any known peptidase domain, it seems other proteases are

419

involved in the substrate degradation. Previously it is reported that a large portion of

420

ClpL is membrane associated suggesting that ClpL may co-operate with

421

membrane-associated proteases (Tran et al., 2011).

422

S. mutans encodes two major membrane-associated stress related proteases, FtsH

423

and HtpX.

In E. coli and B. subtilis, FtsH is induced by heat- and osmotic shocks

424

([43]; [44]).

While FtsH is not essential for growth in species such as B. subtilis,

425

Corynebacterium glutamicum, and Lactobacillus plantarum [44,45]; [46]; FtsH is

426

essential is for some bacteria including E. coli and Lactococcus lactis ([47]; [48]).

427

Our repeated attempts to inactivate ftsH in S. mutans were unsuccessful, suggesting

428

that FtsH is also essential in this organism.

429

is a zinc-dependent mettaloprotease [49]) and is very poorly characterized.

430

we inactivated htpX in S. mutans we found that HtpX was not involved in the

431

degradation of CtsR (data not shown). Thus, additional bioinformatic and biochemical

432

analyses are required to identify the protease involved in CtsR degradation.

Genome analysis indicates that

When

433 434

It appears that CtsR is not the only substrate that is recognized by ClpL. Several

435

proteins, especially some high molecular weight proteins, were differentially

436

expressed in the clpL-deficient strain as compared to its parental strain (data not

437

shown).

438

increased susceptibility to penicillin-induced lysis in S. pneumoniae [31]. The reason

Recently, it has been demonstrated that a knockout mutant of clpL displays

22

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The other stress-related protease, HtpX,

439

for this increased susceptibility is due to reduced PBP2x protein, which is required for

440

cell-wall biosynthesis. That study proposes that ClpL has two functions: “stabilize and

441

reactivate” PBP2x under stressful conditions and facilitate translocation of PBP2x to

442

the cell wall, by some unknown mechanism, which leads to wall thickening and

443

penicillin resistance [31].

444

since we observed increased bacitracin sensitivity in the clpL-deficient strain (data not

445

shown).

446

such as penicillin and cefixime have similar effect on the clpL mutant strain, which

447

displayed increased susceptibility towards these antibiotics.

ClpL probably plays a similar role in S. mutans as well

448 449

Although primary sequence analysis indicates that ClpL is analogous to ClpB protein,

450

a molecular chaperone that is present in both gram-negative and gram-positive

451

bacteria, some important differences also exist between these two proteins.

452

Compared to ClpL, ClpB possesses two additional domains, the N and M domain.

453

Despite the wealth of biochemical and structural data the exact functions of these

454

domains are not fully understood. The N domain of ClpB appears to be dispensable

455

for its oligomerization ability and in vivo chaperone activity [42,50], but it can increase

456

the interaction with protein aggregates [51,52]. On the other hand, the M domain,

457

consisting of four α helices, is very specific for ClpB function and forms a large

458

coiled-coil structure that protrudes from the ClpB hexameric ring [18]. The conserved

459

helix 3 of the M domain is particularly required for the DnaK-dependent shuffling of

460

aggregated proteins and not for soluble denatured substrates [53]. Mutations of this

23

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Furthermore, we also found that several other cell wall damaging agents

domain result in a loss of DnaK/J-GrpE-dependent disaggregation activity while

462

retaining the DnaK/J-GrpE-independent functions of ClpB [42,53,54]. Since the M

463

domain is absent in ClpL protein, we speculate that ClpL does not cooperate with

464

DnaK system and therefore is not able to disaggregate preformed protein aggregates

465

and our observation that ClpL alone was not capable of resolubilizing preformed CtsR

466

aggregates is consistent with this notion.

467

interacts with the target substrates before the formation of protein aggregates and

468

helps to refold the substrates.

469

hexamer in solution [23,42,55].

470

domains are necessary for ClpL oligomerization as demonstrated by our bacterial

471

two-hybrid assays. Our data also indicated that the C-terminal D2-small domain is

472

essential in preventing CtsR aggregation in vivo.

Instead we speculate that ClpL primarily

Our data indicated that ClpL, like ClpB, is present as However, unlike in ClpB, both N- and C- terminal

473 474

In conclusion, we provide evidences that ClpL functions as a chaperone to fold CtsR

475

repressor, an important endogenous protein, and prevents the formation of

476

deleterious protein aggregates during ambient growth condition.

477

chaperone activity is not restricted to stress induced conditions.

478

ClpL plays a critical role in long-term survival [7] is because ClpL overall contributes to

479

the cellular homeostasis by preventing accumulation of protein aggregates. Our in

480

vitro assays suggest that other protein co-factors are not required for this folding

481

activity.

482

during in vivo folding and/or degradation process. Because ClpL plays an important

Therefore, the ClpL The reason that

However, we cannot rule out the possible involvement of other proteins

24

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461

483

role in various cellular functions, detailed biochemical and structural characterizations

484

are necessary to understand the molecular mechanism of this novel chaperone.

485

This will further shed light on how gram-positive bacteria maintain the cellular

486

homeostasis and respond to various stresses.

487

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488 489 490

25

491 492

ACKNOWLEDGEMENTS This work was supported in part by a NIDCR grant (DE021664) to IB.

493 494 495

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26

496

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30

Figure Legends

634

Fig. 1. His-CtsR is accumulated in ΔclpL but mainly as inactive form.

635

A. Western blot analysis of the cell lysates showing the amount of His-CtsR protein

636

in wild-type (UA159) and clpL-deficient strain (IBSJ3) in either soluble fraction (S)

637

or pellet fraction (P). The blot was probed with the anti-His antibody. The same

638

membrane was re-probed with the anti-SmuEno antibody to verify the amount of

639

endogenous enolase, which served as an internal loading control.

640

B. The protein amount of His-HrcA in wild-type and ΔclpL strain in either soluble

641

fraction (S) or pellet fraction (P) was also shown by western blot analysis. The

642

same membrane was also re-probed with the anti-SmuEno antibody.

643

C. Expression of PclpP-gusA fusion in ΔclpL (IBSJ9) compared with wild-type

644

(IBS514). GusA activity was assayed from the PclpP-gusA reporter construct to

645

measure the CtsR repressor activity. The values shown are MU of GusA activity.

646

(n=8 samples/group, p