Escherichia coli cold-shock gene profiles in response to over-expression/deletion of CsdA, RNase R and PNPase
and relevance to low-temperature RNA metabolism

Authors


  • Communicated by: Hiroji Aiba

Correspondence: phadtasa@umdnj.edu

Abstract

Cold-shock response is elicited by the transfer of exponentially growing cells from their optimum temperature to a significantly lower growth temperature and is characterized by the induction of several cold-shock proteins. These proteins, which presumably possess a variety of different activities, are critical for survival and continued growth at low temperature. One of the main consequences of cold shock is stabilization of the secondary structures in nucleic acids leading to hindrance of RNA degradation. Cold-shock proteins, such as RNA helicase CsdA, and 3′-5′ processing exoribonucleases, such as PNPase and RNase R, are presumably involved in facilitating the RNA metabolism at low temperature. As a step toward elucidating the individual contributions of these proteins to low-temperature RNA metabolism, the global transcript profiles of cells lacking CsdA, RNase R and PNPase proteins as well as cells individually over-expressing these proteins as compared to the wild-type cells were analyzed at 15 °C. The analysis showed distinct sets of genes, which are possible targets of each of these proteins. This analysis will help further our understanding of the low-temperature RNA metabolism.

Introduction

Cold-shock response is elicited by the transfer of exponentially growing cells from their optimum temperature to a significantly lower growth temperature and is characterized by the induction of several cold-shock proteins. As cold shock leads to several cellular processes being modified, slowed, or inhibited, these proteins, which presumably possess a variety of different activities prove to be critical for survival and continued growth of the cell at low temperature. NusA (Friedman et al. 1984), IF2 (Gualerzi & Pon 1990), RbfA (Dammel & Noller 1995), CspA (Goldstein et al. 1990) and its homologues such as CspB (Lee et al. 1994), CspG (Nakashima et al. 1996), CspI (Wang et al. 1999), CsdA (Toone et al. 1991; Turner et al. 2007) and PNPase (Reiner 1969a,b; Donovan & Kushner 1986) are among the various major cold-shock proteins produced in Escherichia coli after transfer of the cells from 37 to 15 °C (Jones et al. 1987).

Cold-shock response is well studied from various bacteria (Graumann & Marahiel 1999; Lopez et al. 2001; Balhesteros et al. 2010) to higher organisms (Bradley et al. 2007). One of the main consequences of cold shock is stabilization of the secondary structures in nucleic acids (Rajkowitsch et al. 2007; Phadtare & Severinov 2010; Phadtare 2011) leading to hindrance of (i) transcription and translation and (ii) RNA degradation. The role of CspA and its homologues as RNA chaperones that act as transcription antiterminators and essentiality of this activity for the cold acclimation of cells has been well studied and the Csp-responsive, promoter-proximal sequences that can block the transcript elongation have been identified in several target genes of Csp homologues (Jiang et al. 1997; Bae et al. 2000; Phadtare et al. 2002, 2006; Phadtare & Inouye 2004). However, cold-shock proteins such as RNA helicase, CsdA and 3′-5′ processing exoribonucleases such as PNPase and RNase R are presumably involved in facilitating the RNA metabolism at low temperature. In the present study, we will focus on these three proteins in the context of the cellular cold-shock response.

CsdA is a highly conserved, DEAD-box RNA helicase (Linder et al. 1989) and is essential only at low temperature (Jones et al. 1996; Charollais et al. 2004). It has been suggested to be involved in the biogenesis of the small ribosomal subunits (Toone et al. 1991; Moll et al. 2002) and the 50S ribosomal subunits (Charollais et al. 2004), promotion of translation initiation of structured mRNAs (Lu et al. 1999), low-temperature riboregulation of RpoS mRNA (Resch et al. 2010) and stabilization and degradation of mRNAs (Tamura et al. 2012; Khemici et al. 2004; Prud'homme-Genereux et al. 2004; Yamanaka & Inouye 2001). Its role in mRNA decay and ribosome biogenesis has been studied in detail. Interestingly, the unwinding (helicase) activity of CsdA may be important for both of these functions. It was suggested that CsdA may help 50S assembly by modulating RNA or RNP (ribonucleoprotein) structures, and its unwinding activity may be required to facilitate structural transitions within the RNA and may also allow proper binding of ribosomal protein(s) (Iost & Dreyfus 2006). It may alternatively prevent and/or resolve misfolding, which may provide assistance to rRNA to reach its active conformation. Our studies showed that CsdA-mediated mRNA decay may be critical during cold shock, and the helicase activity of CsdA is crucial for promoting degradation of mRNAs stabilized at low temperature (Awano et al. 2007). We showed that a target mRNA was significantly stabilized in the csdA null mutant cells at 15 °C, and this effect was counteracted by over-expression of wild-type CsdA protein but not by a helicase-deficient mutant of CsdA. Furthermore, our in vivo genetic screening of an Ecoli csdA null mutant strain and results from other research groups showed that RhlE, another DEAD-box RNA helicase, can complement the cold sensitivity of the csdA null mutant strain (Iost & Dreyfus 2006; Awano et al. 2007; Jain 2008). We also observed that two cold-shock proteins, CspA and RNase R can also substitute for CsdA at low temperature, albeit to a somewhat weaker degree. It is also interesting to note that combined absence of CsdA and RNase R results in increased sensitivity of the cells to moderate temperature downshifts.

RNase R, PNPase and RNase II are the three major 3′-to-5′ processing exoribonucleases in Ecoli. These enzymes are primarily involved in RNA metabolism in Ecoli. PNPase (Jones et al. 1987; Zangrossi et al. 2000) and RNase R (Cairrao et al. 2003) are induced by cold shock and are suggested to be the universal degraders of structured RNA in vivo (Li et al. 2002; Cheng & Deutscher 2003). Escherichia coli PNPase (polynucleotide phosphorylase) is essential for growth at low temperatures (Luttinger et al. 1996; Piazza et al. 1996). However, its exact role in this essential function is not fully elucidated. In addition to promoting the processive degradation of RNA, PNPase is also responsible for residual RNA tailing observed in E. coli mutants devoid of the main polyadenylating enzyme PAP I (Mohanty & Kushner 2000a,b, 2003).

We have made several interesting observations during further studies on these proteins, (i) of these three exoribonucleases, only RNase R can substitute for CsdA at low temperature (Awano et al. 2007), (ii) upon in vivo domain analysis of RNase R, we showed that it also has helicase activity, and this activity is essential to substitute for the cold-shock function of CsdA. RNase R possesses ribonuclease and helicase activities, which are distinct from each other as the mutant RNase R proteins lacking the ribonuclease activity retained their ability to substitute for CsdA. It was observed that the CSD2 domain of RNase R is not essential for its ribonuclease activity; however, it is important for its helicase activity (Awano et al. 2010), (iii) ribonuclease activity of PNPase is critical at low temperature (Awano et al. 2008); however, in spite of its ability to act as the universal degrader of the structured RNAs in vivo (Li et al. 2002; Cheng & Deutscher 2003), it cannot substitute for CsdA at low temperature (Awano et al. 2007) and (iv) RNase II and RNase R belong to the RNR family and exhibit approximately 60% similarity in their secondary structures, but RNase II can substitute for PNPase and not CsdA, whereas RNase R can substitute for CsdA, but not PNPase at low temperature (Awano et al. 2007, 2008).

These observations suggest that (i) CsdA and RNase R share common target mRNAs, whose degradation they facilitate by virtue of their unwinding activity. Note that the target mRNA, which was significantly stabilized in the csdA null mutant cells at 15 °C was destabilized by over-expression of CsdA as well as RNase R (Awano et al. 2007). (ii) Some of these targets, which are apparently important for the cold acclimation of cells, cannot be degraded by PNPase leading to the lack of suppression of the cold sensitivity of the csdA null mutant strain by the PNPase over-expression. (iii) There exists a distinct set of target mRNAs degraded by PNPase, which may be relevant to the cold-shock response of the cells. However, these mRNAs are not degraded by RNase R which would explain the lack of suppression of the cold sensitivity of the Δpnp strain by RNase R. Thus, it can be hypothesized that these proteins have different substrate specificities. Previously, it was reported that although CsdA does not have strong specificity for RNA substrates in vitro, it may have substrate preferences in vivo (Iost & Dreyfus 2006). The helicase activity of RNase R is independent of its ribonuclease activity; thus, the RNA substrates for these activities may be different or it may share a common subset of mRNAs, which it can both unwind and degrade. Although RNase R may have some preference for RNA substrates, it can degrade (Awano et al. 2008); no data are available at present as to the preferred targets for its unwinding activity. A comparative DNA microarray analysis of the targets of these three proteins at 15 °C should provide clues to elucidate how RNA metabolism is mediated at low temperature. Thus, in the present study, DNA microarray analysis of the effect of deletion and over-expression of CsdA, RNase R and PNPase at 15 °C as compared to the wild-type cells was carried out.

To get a comprehensive picture of the changes occurring at low temperature, these data were also compared with our published DNA microarray data (Phadtare & Inouye 2004) profiling the genes which are affected by the temperature downshift in the wild-type cells as well as cold-sensitive cells bearing deletions of the four csp genes.

Results and discussion

The goal of this work is to identify all of the E. coli open reading frames (ORF) whose mRNAs undergo a significant change in abundance in response to changes in the levels of CsdA, RNase R and PNPase in the cells at 1 h upon cold shock (15 °C). The following conditions were investigated: (i) cells lacking either csdA, rnr or pnp and growing at 15 °C and (ii) wild-type cells growing at 15 °C and overproducing CsdA, RNase R or PNPase from plasmids.

Total cellular RNA was isolated and used to generate probes that were hybridized to DNA arrays, and the results were quantified as described in the 'Experimental procedures'. Each microarray experiment was independently repeated two times. The cell density of the control and test cells for each set was the same; thus, the observed changes in mRNA levels were not because of the differences in cell densities. For over-expression of respective proteins, appropriate isopropyl β-D thiogalactopyranoside (IPTG)-inducible pINIII vector-based plasmids were used. Cells carrying the pINIII vector served as control for these sets, whereas the wild-type E. coli JM83 cells were used as control for the deletions strains. The csdA null mutant, Δrnr and Δpnp strains and plasmids for over-expressing these proteins have been reported previously (Awano et al. 2007, 2008, 2010); these were tested in vivo before the DNA microarray analysis as shown in Fig. 1. Consistent with the previous reports (Luttinger et al. 1996; Awano et al. 2007, 2008), the wild-type and the three deletion strains were able to grow at 37 °C (Fig. 1A), and the wild-type and the Δrnr strains are also able to grow at 15 °C, whereas the csdA null mutant and Δpnp strains were not (Fig. 1B). Note that, Δrnr strain by itself is not cold-sensitive, whereas the double mutant (csdA::kanΔrnr) strain exhibits cold sensitivity even at 20 °C (Awano et al. 2007). The cold sensitivity of the csdA null mutant strain was suppressed by the pINIII-based over-expression of CsdA and RNase R and that of the Δpnp strain was suppressed by over-expression of PNPase (Fig. 1C). Consistent with previous reports, the cold sensitivity of the csdA null mutant and Δpnp strains was not suppressed by the over-expression of PNPase and RNase R, respectively (Awano et al. 2007, 2008). As expected, the deletion of the respective genes was apparent in the DNA microarray analysis. The DNA microarray data showed increase in the level of mRNAs for these three genes in the cells carrying the respective pINIII constructs; 1.8, 3.84 and 1.8-fold increase was seen for csdA, rnr and pnp, respectively.

Figure 1.

Cold sensitivity of the csdA null mutant and Δpnp strains used for the study and suppression of their cold sensitivity by over-expression of respective proteins. The E. coli wild-type, csdA null mutant (csdA::kan), Δpnp and Δrnr cells were streaked on LB plates and grown at 37 °C (A) or 15 °C (B). The Ecoli wild-type cells transformed with pINIII plasmid as control and the csdA null mutant or Δpnp cells transformed with pINIII plasmid alone as control, or expressing CsdA or RNase R or PNPase were streaked on LB plates containing ampicillin (50 μg/mL) and were grown at 15 °C (C). Results of the plates incubated at 37 °C for 24 h and 15 °C for 120 h are presented.

Supporting information Tables list the genes which show 1.5-fold difference at 15 °C in response to the deletion (Table S1 in Supporting Information) and over-expression (Table S2 in Supporting Information) of csdA, rnr or pnp as compared to the wild-type cells treated similarly. Note that, genes which show 1.5-fold change (increase or decrease) in at least one of the test conditions are included in these Tables. As described in the 'Experimental procedures' section, 1.5-fold change is statistically significant. Positive values suggest up-regulation and negative values suggest down-regulation. A subgroup of Tables was then created by tabulating genes which show more significant change (equal to or more than 2-fold) with either of the three test conditions and these were categorized as: genes that are down-regulated in all three deletions strains (Table 1), up-regulated in all three deletions strains (Table 2) and differentially affected (up-regulated in one test condition and down-regulated in the other) in the three deletion strains (Tables 3 and 4), and genes that are down-regulated in response to all the three over-expressions (Table 5), up-regulated in response to all three over-expressions (Table 6) and differentially affected in response to the three over-expressions (Table 7). In each Table, genes are furthermore categorized based on their cellular role. Please note that, genes encoding hypothetical, putative or predicted proteins are not included in the Tables 1-7. For the analysis of the genes which are affected by the three deletions, it was also checked if this effect was counteracted by over-expression of the respective proteins by comparing with the data presented in Tables 5-7 or the Table S2 in Supporting Information. Similarly, while analyzing the genes that are affected by over-expression of csdA/rnr/pnp, it was also analyzed whether this effect was reversed by the deletion of respective genes. As mentioned above, comparisons were also made with our published data (Phadtare & Inouye 2004) with respect to the genes reported to be important for cold-shock response and acclimation of cells. Results obtained by the DNA microarray analysis were confirmed by qRT-PCR analysis of few selected genes as shown in Fig. 2. These genes were randomly selected from all the Tables, RNAs were independently isolated from the cells for respective conditions and subjected to qRT-PCR as described in detail in the 'Experimental procedures'. The assays were carried out in triplicate. Each of the genes selected for the qRT-PCR analysis showed similar changes, that is, up- or down-regulation, in their levels as seen in the DNA microarray confirming the validity of the data obtained. For example, the qRT-PCR analysis showed that cheA is down-regulated (0.48) in the csdA null mutant strain and up-regulated (7.97) in the Δpnp strain. The DNA microarray data (Table 3) shows respective changes (−2.42 in csdA null mutant strain and 2.99 in the Δpnp strain) for cheA.

Figure 2.

Effect of over-expression and deletion of CsdA, RNase R and PNPase on the levels of selected mRNAs. Results of qRT-PCR analysis carried out for nine genes are shown. Genes to be tested for qRT-PCR were randomly selected from all the data tables (Tables 1-6) presented in the text and analysis was carried out as described in the 'Experimental procedures'. For the level of genes in the deletion cells, their respective level in wild-type is normalized to 1.0; for the level of genes in the cells over-expressing either CsdA, RNase R or PNPase, their respective level in wild-type cells carrying the vector alone is normalized to 1.0. The actual values along with the standard deviation values are shown in Table S3 in Supporting Information.

Table 1. Genes down-regulated in all three deletion strains
GeneGene product and/or function

csdA::kan/wt

15 °C, 1 h

Δrnr/wt

15 °C, 1 h

Δpnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
cysA Sulfate/thiosulfate transporter subunit−4.08−1.22−1.11
cysP Thiosulfate transporter subunit−8.62−2.82−1.26
cysU Sulfate/thiosulfate transporter subunit−4.33−2.00−1.15
cysW Sulfate/thiosulfate transporter subunit−4.51−1.80−1.50
gltP Glutamate/aspartate:proton symporter−2.19−2.06−1.58
malE Maltose ABC transporter periplasmic protein−2.31−1.98−1.91
zntA Zinc/cadmium/mercury/lead-transporting ATPase−2.72−1.17−1.20
Genes involved in cell metabolism
aceA Isocitrate lyase−1.98−1.18−2.37
aceB Malate synthase−1.82−1.05−2.06
aceK Isocitrate dehydrogenase kinase/phosphatase−2.45−1.35−2.37
acs Acetyl-CoA synthetase−2.12−2.01−2.58
cynS Cyanate hydratase−1.75−2.01−3.57
cysD Sulfate adenylyltransferase−9.79−2.99−1.50
cysH 3-phosphoadenosine 5-phosphosulfate reductase−4.11−1.49−1.45
cysI Sulfite reductase−3.94−1.55−1.48
cysJ Sulfite reductase subunit alpha−4.99−2.44−1.65
cysN Sulfate adenylyltransferase−3.50−1.25−1.20
lysA Diaminopimelate decarboxylase−6.63−2.71−2.73
nrdA Ribonucleotide-diphosphate reductase subunit alpha−2.23−1.76−1.07
nrdB Ribonucleotide-diphosphate reductase 1−2.23−1.94−1.35
uxaC Uronate isomerase−1.04−1.01−2.17
Genes encoding proteins with diverse functions
bdm Biofilm-dependent modulation protein−1.03−1.07−2.82
dnaJ Chaperone protein DaJ−1.63−1.44−2.20
dnaK Molecular chaperone DnaK−1.59−1.81−2.18
htpG Heat shock protein 90−1.66−1.48−2.23
sppA Protease 4−1.02−1.41−2.03
Table 2. Genes up-regulated in all three deletion strains
GeneGene product and/or function

csdA::kan/wt

15 °C, 1 h

Δrnr/wt

15 °C, 1 h

Δpnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
bglF Fused beta-glucoside-specific PTS enzymes1.071.532.59
flgA Flagellar basal body P-ring biosynthesis protein FlgA1.451.312.45
flgB Flagellar basal body rod protein FlgB2.071.562.70
flgD Flagellar hook assembly protein1.651.522.54
flgE Flagellar hook protein FlgE1.521.502.24
flgG Flagellar basal body rod protein FlgG1.681.672.28
flgH Flagellar basal body l-ring protein1.751.912.56
flgJ Flagellar rod assembly protein/muramidase FlgJ1.501.952.81
flgL Flagellar hook-associated protein FlgL1.101.023.59
fliA Flagellar biosynthesis sigma factor1.021.119.16
fliG Flagellar motor switch protein G1.322.562.09
fliJ Flagellar biosynthesis chaperone1.603.222.77
fliK Flagellar hook-length control protein1.652.892.44
fliL Flagellar basal body-associated protein FliL1.912.714.05
fliN Flagellar motor switch protein FliN1.602.302.18
fliO Flagellar biosynthesis protein FliO1.592.322.33
fliS Flagellar protein FliS1.091.193.20
modC Molybdate transporter ATP-binding protein1.121.182.38
modF Fused molybdate transporter subunits of ABC superfamilys1.161.336.55
pbpC Penicillin-binding protein 1C1.672.121.01
Genes involved in cell metabolism
aslA Arylsulfatase1.031.202.48
bglG Transcriptional antiterminator BglG1.011.202.48
ilvG Acetolactate synthase 2 catalytic subunit1.092.271.46
leuA 2-isopropylmalate synthase1.612.211.01
ndh NADH dehydrogenase3.311.282.33
pta Phosphate acetyltransferase2.161.281.61
Genes encoding proteins with diverse functions
csgD DNA-binding transcriptional regulator CsgD1.671.182.26
nth Endonuclease III1.332.271.18
rhsC rhsC protein1.001.143.35
Table 3. Genes showing similar response to only csdA and rnr deletions
GeneGene product and/or function

csdA::kan/wt

15 °C, 1 h

Δrnr/wt

15 °C, 1 h

Δpnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
arsB Arsenite/antimonite transporter−3.62−1.531.07
cheA Chemotaxis protein CheA−2.42−1.342.99
cheB Chemotaxis-specific methylesterase−2.25−1.441.86
cheW Purine-binding chemotaxis protein−2.25−1.292.99
cheY Chemotaxis regulatory protein CheY−2.32−1.531.51
fhuB Iron-hydroxamate transporter permease subunit1.122.06−1.43
fimB Tyrosine recombinase−2.34−1.022.08
flgM Anti-sigma28 factor FlgM−1.14−1.303.86
flgN Flagella synthesis protein FlgN−1.24−1.083.57
fliC Flagellar filament structural protein (flagellin)−1.53−1.192.75
hlyE Hemolysin E−1.16−1.093.31
kch Voltage-gated potassium channel−1.04−2.272.63
motA Flagellar motor protein MotA−2.83−1.373.83
motB Flagellar motor protein MotB−2.38−1.373.11
ompN Outer membrane pore protein N−1.08−1.102.38
sbp Sulfate transporter subunit−2.47−1.771.18
sulA SOS cell division inhibitor−1.61−1.492.07
tap Methyl-accepting protein IV−1.77−1.232.56
tar Methyl-accepting chemotaxis protein II−2.01−1.444.52
tsr Methyl-accepting chemotaxis protein I−2.13−1.721.98
Genes involved in cell metabolism
cdh CDP-diacylglycerol pyrophosphatase−2.16−1.531.07
cysC Adenylylsulfate kinase−3.68−1.441.03
hybD Hydrogenase 2 maturation endopeptidase−1.35−1.632.20
leuA 2-isopropylmalate synthase1.602.10−1.25
leuB 3-isopropylmalate dehydrogenase1.572.00−1.27
soxS DNA-binding transcriptional regulator SoxS−1.23−2.582.60
trpD Bifunctional glutamine amidotransferase1.211.18−2.10
trpE Component I of anthranilate synthase1.041.04−2.18
ybaS Glutaminase2.101.56−1.60
Genes encoding proteins with diverse functions
appY DNA-binding transcriptional activator−1.18−1.052.14
arsC Arsenate reductase−2.37−1.591.19
arsR DNA-binding transcriptional repressor−3.99−1.781.21
cspH Cold-shock-like protein1.451.402.62
dinD DNA damage-inducible protein−1.37−1.282.00
dinI DNA damage-inducible protein I−1.23−1.592.40
recN Recombination and repair protein−1.12−1.272.95
recX Recombination regulator RecX−1.42−1.383.76
rnr Exoribonuclease R///exoribonuclease R−1.13−185.95(NA)1.09
stpA DNA-binding protein−1.24−1.222.33
umuC DNA polymerase V−1.99−1.464.58
umuD DNA polymerase V subunit UmuD−2.44−1.84.43
yebG DNA damage-inducible protein YebG−1.25−1.372.16
Table 4. Genes showing different response to csdA and rnr deletions
GeneGene product and/or function

csdA::kan/wt

15 °C, 1 h

Δrnr/wt

15 °C, 1 h

Δpnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
amtB Ammonium transporter−1.012.051.29
arpA Ankyrin repeat protein−4.911.04−1.26
cutC Copper homeostasis protein CutC−2.571.041.08
fecA Ferric citrate outer membrane transporter−1.212.42−1.24
fecB Iron-dicitrate transporter subunit−1.092.62−1.50
fecC Iron-dicitrate transporter subunit1.243.12−1.79
fecD Iron-dicitrate transporter subunit1.333.18−1.36
fecE Iron-dicitrate transporter subunit1.322.82−1.06
fecI RNA polymerase, Sigma 19 factor−1.221.92−1.12
fecR Transmembrane signal transducer for ferric citrate transport−1.182.66−1.10
fhuD Iron-hydroxamate transporter substrate-binding subunit−1.022.18−1.58
fimC FimC−2.411.801.53
fimD FimD−2.473.151.83
fimF FimF−2.493.371.88
fimG FimG−2.183.841.93
fimH Minor fimbrial subunit−1.893.391.75
fimI FimI−2.191.551.41
flgK Flagellar hook-associated protein FlgK−1.021.264.47
flhC Transcriptional activator FlhC−1.211.472.43
flhD Transcriptional activator FlhD−1.391.462.24
fliD Flagellar capping protein−1.081.173.37
gntP Fructuronate transporter−1.022.691.97
hokA Toxic polypeptide−3.141.041.69
nuoM NADH dehydrogenase subunit M1.08−1.13−2.02
pstS PstS−1.271.042.33
rbsA d-ribose transporter ATP-binding protein−2.791.82−1.25
rbsB d-ribose transporter subunit RbsB−2.101.09−1.60
rbsC Ribose ABC transporter permease protein−2.281.27−1.59
thiQ Thiamine transporter ATP-binding subunit−1.292.101.26
tnaB Tryptophan permease TnaB−1.131.162.18
ygiZ Conserved inner membrane protein−1.121.152.27
Genes involved in cell metabolism
eutB Ethanolamine ammonia-lyase−1.071.452.35
folP Dihydropteroate synthase−1.781.14−3.82
glmM Phosphoglucosamine mutase−1.631.16−2.23
glnA Glutamine synthetase−1.052.081.05
glnG Nitrogen regulation protein NR(I)−1.683.25−1.06
glnL Nitrogen regulation protein NR(II)−1.292.56−1.20
glnK Nitrogen regulatory protein P-II 2−1.423.221.62
glpC sn-glycerol-3-phosphate dehydrogenase (anaerobic)−1.152.101.10
glpK Glycerol kinase−2.181.00−1.54
hisD Histidinol dehydrogenase1.14−1.10−2.17
ilvB Acetolactate synthase catalytic subunit−2.231.48−1.27
ilvG Acetolactate synthase 2 catalytic subunit−1.052.361.49
ilvN Acetolactate synthase 1 regulatory subunit−2.111.57−1.53
malT Transcriptional regulator MalT−2.002.15−1.79
purC Phosphoribosylaminoimidazole-succinocarboxamide synthase−1.382.021.10
purF Amidophosphoribosyltransferase−1.392.351.52
purL Phosphoribosylformyl-glycineamide synthetase−1.122.451.16
Genes encoding proteins with diverse functions
cdaR DNA-binding transcriptional activator−2.991.69−1.04
deaD CsdA−2.001.291.49
degP (htrA) Serine endoprotease−3.421.25−1.14
dhaK Dihydroxyacetone kinase−2.111.2−2.26
dinB (dinP) DNA polymerase IV−1.131.123.08
pnp Polynucleotide phosphorylase−1.011.16257.64(NA)
sfa Cold-shock gene−2.721.121.24
zntR Zinc-responsive transcriptional regulator1.05−1.042.10
Table 5. Genes showing down-regulation in response to overproduction of all three proteins
GeneGene product and/or function

↑csdA/wt

15 °C, 1 h

↑rnr/wt

15 °C, 1 h

↑pnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
aroP Aromatic amino acid transporter−1.27−2.11−1.25
celA N,N'-diacetylchitobiose-specific PTS transporter subunit IIB−2.61−1.27−1.46
celB N,N'-diacetylchitobiose-specific PTS transporter subunit IIC−2.18−1.26−1.17
celC N,N'-diacetylchitobiose-specific PTS transporter subunit IIA−2.29−1.24−1.28
cysP Thiosulfate transporter subunit−2.58−1.06−1.66
dsbG Periplasmic disulfide isomerase/thiol-disulfide oxidase−2.16−1.49−1.10
entD Phosphopantetheinyltransferase−1.29−2.37−1.32
fic Cell filamentation protein Fic−2.35−1.32−1.35
fimA FimA−2.16−1.06−2.12
fhuE Ferric-rhodotorulic acid outer membrane transporter−4.00−2.61−1.23
gltF Periplasmic protein−2.74−1.25−1.10
gntX Gluconate periplasmic binding protein−2.23−1.08−1.14
hdeD Acid-resistance membrane protein−7.81−2.31−2.52
mdtE Multidrug efflux system protein MdtE−6.06−1.54−1.65
mdtF Putative transport system permease protein−3.09−1.42−1.35
mntH Manganese transport protein MntH−2.25−4.18−1.82
nlpA Cytoplasmic membrane lipoprotein-28−2.68−1.19−1.31
nupC Nucleoside permease nupC−2.42−1.66−1.14
ompC Outer membrane porin protein C−2.47−1.09−3.56
papH PapH protein///PapH protein−2.07−1.30−2.79
pntA NAD(P) transhydrogenase subunit alpha−2.07−1.42−1.35
shiA Shikimate transporter−2.76−2.57−2.20
slp Slp−6.89−1.72−1.66
sufC Cysteine desulfurase ATPase component−3.07−2.79−2.28
tyrP Tyrosine transporter−2.09−2.62−1.85
yddR ABC transporter permease−2.17−1.15−1.13
Genes involved in cell metabolism
aceA Isocitrate lyase−1.66−1.10−2.85
argE Acetylornithine deacetylase−2.46−1.32−1.09
aroF Phospho-2-dehydro-3-deoxyheptonate aldolase−1.70−2.71−2.58
aspC Aromatic amino acid aminotransferase−2.08−1.31−1.18
cbl Transcriptional regulator Cbl -cysteine biosynthesis−1.81−1.1−2.13
celD DNA-binding transcriptional regulator ChbR−2.14−1.07−1.13
chbF Cryptic phospho-beta-glucosidase, NAD(P)-binding−2.08−1.11−1.05
citC Citrate lyase synthetase (citrate (pro-3S)-lyase ligase−2.98−1.57−1.69
deoD Purine-nucleoside phosphorylase−2.03−1.05−1.18
dld d-lactate dehydrogenase−2.38−1.42−1.31
dos cAMP phosphodiesterase−3.34−1.36−2.35
gadA Glutamate decarboxylase isozyme−4.82−2.55−3.36
gadB Glutamate decarboxylase isozyme−19.51−4.16−4.05
ghrB 2-hydroxyacid dehydrogenase−2.04−1.45−1.47
glgX Glycogen debranching enzyme−1.73−1.90−2.11
gltD Glutamate synthase−1.22−2.43−1.65
gmhB d,d-heptose 1,7-bisphosphate phosphatase−2.05−1.12−1.21
gpmA Phosphoglyceromutase−2.09−1.41−1.29
grxB Glutaredoxin 2−2.20−1.25−1.26
gst Glutathionine S-transferase−2.03−1.26−1.29
hisL his operon leader peptide−2.04−1.13−1.09
leuL leu operon leader peptide−2.12−1.18−1.20
manA Mannose-6-phosphate isomerase−2.15−1.07−1.12
msrA Methionine sulfoxide reductase A−3.03−1.72−1.83
nadE NAD synthetase−2.22−1.93−1.71
nrdH Glutaredoxin-like protein−1.39−2.2−2.47
otsB Trehalose-6-phosphate phosphatase−1.09−2.04−1.70
pgl 6-phosphogluconolactonase−1.65−2.00−1.48
pptA 4-oxalocrotonate tautomerase−4.07−1.77−1.41
rimL Ribosomal protein-L7/L12-serine acetyltransferase−2.09−1.56−1.12
rpiA Ribose-5-phosphate isomerase A−2.61−1.08−1.80
sufA Fe-S cluster assembly protein−3.71−2.50−1.90
sufB Cysteine desulfurase activator complex subunit SufB−3.06−2.76−1.99
sufD Cysteine desulfurase activator complex subunit SufD−3.19−3.05−1.93
sufE Cysteine desufuration protein SufE−4.41−4.73−2.81
sufS Bifunctional cysteine desulfurase/selenocysteine lyase−3.31−3.23−2.09
tyrA Bifunctional chorismate mutase/prephenate dehydrogenase−2.14−3.62−4.17
tyrB Aromatic amino acid aminotransferase−3.56−2.10−1.90
ybaS Glutaminase−2.91−1.65−1.93
ybiL Catecholate siderophore receptor Fiu−2.36−1.57−1.03
Genes encoding proteins with diverse functions
bdm Biofilm-dependent modulation protein−2.06−1.21−2.76
cspC Cold-shock-like protein CspC−2.11−1.22−1.24
hdeA Acid-resistance protein−26.28−1.87−2.06
hdeB Acid-resistance protein−25.40−2.14−2.46
nrdI Ribonucleotide reductase stimulatory protein−1.36−2.85−2.59
raiA Translation inhibitor protein RaiA−2.85−1.11−2.08
uspC Universal stress protein UspC−1.45−1.34−2.70
yfaY Competence damage-inducible protein A−2.00−1.40−1.25
Table 6. Genes showing up-regulation in response to overproduction of all three proteins
GeneGene product and/or function

↑csdA/wt

15 °C, 1 h

↑rnr/wt

15 °C, 1 h

↑pnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
aer Aerotaxis sensor receptor, flavoprotein2.771.341.87
ais Protein induced by aluminum///Ais protein2.851.471.39
amiB N-acetylmuramoyl-l-alanine amidase II2.011.161.66
ansP l-asparagine permease1.392.101.82
apaG ApaG///protein associated with Co2+ and Mg2+ efflux2.361.091.33
arsB Arsenite/antimonite transporter2.021.621.48
bcsB Regulator of cellulose synthase2.291.212.30
ccmG Periplasmic thioredoxin of cytochrome c-type biogenesis2.471.101.30
ccmH heme lyase, CcmH subunit2.091.251.52
cls Cardiolipin synthetase///cardiolipin synthase 13.021.331.67
coaD Phosphopantetheine adenylyltransferase2.291.031.70
cydC Cysteine/glutathione ABC transporter membrane/ATP-binding component2.031.041.30
dctA C4-dicarboxylate transporter DctA1.733.191.11
ddlB d-alanine–d-alanine ligase2.611.011.13
ddpA d-ala-d-a la transporter subunit putative2.331.201.00
ddpC d-ala-d-ala transporter subunit2.041.101.09
ddpF d-ala-d-ala transporter subunit2.641.061.06
emrD Multidrug resistance protein D5.443.114.01
fepC Iron-enterobactin transporter ATP-binding protein2.341.031.49
fepE Ferric enterobactin transport protein FepE2.111.261.27
fepG Iron-enterobactin transporter permease2.301.031.73
ftsA Cell division protein FtsA2.241.061.16
ftsL Cell division protein FtsL2.511.051.20
ftsW Cell division protein FtsW3.701.091.41
gspC General secretion pathway protein C4.412.162.44
gspF Putative general secretion pathway protein F2.301.131.42
gspK General secretory pathway component2.041.021.17
gspM Putative general secretion pathway protein M2.411.211.54
imp Organic solvent tolerance protein2.171.091.09
kch Voltage-gated potassium channel10.393.773.69
kefA Potassium-efflux protein KefA2.051.161.82
lolC Outer membrane-specific lipoprotein transporter subunit LolC2.241.131.23
lolE Outer membrane-specific lipoprotein transporter subunit LolE2.561.061.32
lpxB Tetraacyldisaccharide-1-P synthase///lipid-A-disaccharide synthase2.811.281.40
malG Maltose transporter permease3.111.541.18
metN dl-methionine transporter ATP-binding subunit1.842.222.51
mltA Murein transglycosylase A2.371.061.31
mraY Phospho-N-acetylmuramoyl-pentapeptide transferase2.301.011.29
mrcA Peptidoglycan synthetase2.931.532.13
mrdA Penicillin-binding protein 23.651.441.74
mrdB Cell wall shape-determining protein3.121.411.84
murB UDP-N-acetylenolpyruvoylglucosamine reductase2.011.221.21
murC UDP-N-acetylmuramate–l-alanine ligase2.861.111.23
murD UDP-N-acetylmuramoyl-l-alanyl-d-glutamate synthetase2.681.031.44
murE UDP-N-acetylmuramoylalanyl-d-glutamate–2,6-diaminopimelate ligase2.341.061.14
murF UDP-N-acetylmuramoyl-tripeptide:d-alanyl-d-alanine ligase2.631.101.18
murG N-acetylglucosaminyl transferase3.381.141.32
nfrB Bacteriophage N4 adsorption protein B2.041.602.15
nuoJ NADH dehydrogenase subunit J2.221.732.03
nuoH NADH dehydrogenase subunit H2.991.882.15
nuoK NADH dehydrogenase subunit K2.621.912.24
nuoL NADH dehydrogenase subunit L2.561.722.46
nuoN NADH dehydrogenase subunit N1.911.562.28
ompN Outer membrane protein N precursor3.541.991.63
pheP Phenylalanine transporter1.931.832.12
pqiA Paraquat-inducible protein A3.261.091.27
pqiB Paraquat-inducible protein B2.931.131.30
pspD Peripheral inner membrane phage-shock protein2.311.501.82
pstA Phosphate transporter permease subunit PtsA2.101.231.13
rbsA d-ribose transporter ATP-binding protein23.522.721.03
rcsC Hybrid sensory kinase4.511.091.57
rcsD Phosphotransfer intermediate protein2.101.021.04
sieB Phage superinfection exclusion protein4.071.371.95
ssuA Alkanesulfonate transporter subunit1.056.271.09
ssuC Alkanesulfonate transporter permease subunit1.084.561.30
sulA SOS cell division inhibitor3.472.071.55
tauC Taurine transporter subunit1.063.211.00
tolR Colicin uptake protein TolR2.311.131.49
torS Hybrid sensory histidine kinase2.291.091.45
uhpB Sensory histidine kinase UhpB2.981.182.09
uhpC Regulatory protein UhpC4.151.652.44
zntA Zinc/cadmium/mercury/lead-transporting ATPase2.161.031.64
Genes involved in cell metabolism
ada O6-methylguanine-DNA methyltransferase1.591.403.00
alkB DNA repair system specific for alkylated DNA1.441.252.38
apaH Diadenosine tetraphosphatase2.591.151.45
arsR DNA-binding transcriptional repressor2.041.671.26
bcsA Cellulose synthase, catalytic subunit2.031.161.96
bcsC Cellulose synthase subunit BcsC2.101.061.35
bglF Fused beta-glucoside-specific PTS enzymes2.811.261.45
birA Biotin–protein ligase2.141.171.31
bisZ Biotin sulfoxide reductase 25.081.111.02
carA Glutamine amidotransferase2.231.721.43
carB Carbamoyl phosphate synthase large subunit2.601.931.30
cdd Cytidine deaminase1.222.391.02
cpsB Mannose-1-phosphate guanyltransferase2.121.161.54
cynS Cyanate hydratase5.021.691.59
dacB d-alanyl-d-alanine carboxypeptidase2.231.251.30
dgsA DNA-binding transcriptional repressor2.051.131.10
dkgB 2,5-diketo-d-gluconate reductase B3.612.233.17
dusC tRNA-dihydrouridine synthase C2.361.041.54
dxs 1-deoxy-d-xylulose-5-phosphate synthase2.501.281.44
erpA Iron-sulfur cluster insertion protein ErpA2.151.451.53
eutB Ethanolamine ammonia-lyase heavy chain3.221.221.60
fabB 3-oxoacyl-(acyl carrier protein) synthase I2.081.392.83
fadA 3-ketoacyl-CoA thiolase2.321.651.27
fadB Multifunctional fatty acid oxidation complex subunit alpha2.072.271.03
fadJ Multifunctional fatty acid oxidation complex subunit alpha2.651.681.01
fdhD Formate dehydrogenase accessory protein2.131.031.33
gidA tRNA uridine 5-carboxymethylaminomethyl modification enzyme GidA2.781.221.44
glmS Glucosamine–fructose-6-phosphate aminotransferase2.451.061.34
glpX Fructose 1,6-bisphosphatase II2.241.251.07
gsk Inosine-guanosine kinase2.131.201.42
guaB IMP dehydrogenase2.131.111.11
hemA Glutamyl-tRNA reductase2.461.411.60
hemK N5-glutamine S-adenosyl-l-methionine-dependent methyltransferase2.191.41.74
hlyE Hemolysin E2.811.211.09
hokA Toxic polypeptide, small5.671.431.50
hyaF Hydrogenase-1 operon protein HyaF2.041.081.10
hybD Hydrogenase 2 maturation endopeptidase5.352.591.39
iap Alkaline phosphatase isozyme conversion aminopeptidase3.591.091.50
idnK d-gluconate kinase, thermosensitive2.361.171.25
ileS Isoleucyl-tRNA synthetase2.211.121.27
ilvG Acetolactate synthase 2 catalytic subunit3.941.171.18
ilvM Acetolactate synthase 2 regulatory subunit3.351.321.18
ilvN Acetolactate synthase 1 regulatory subunit2.691.801.03
ispH 4-hydroxy-3-methylbut-2-enyl diphosphate reductase2.431.111.10
kdtA 3-deoxy-d-manno-octulosonic-acid transferase3.191.392.17
kduD 2-deoxy-d-gluconate 3-dehydrogenase2.571.161.13
ksgA Dimethyladenosine transferase2.081.211.21
lacA Galactoside O-acetyltransferase2.951.791.06
lolD Outer membrane-specific lipoprotein transporter subunit2.601.171.40
lrhA Transcriptional regulator LYSR-type3.431.551.91
maa Maltose O-acetyltransferase2.091.331.70
melA Alpha-galactosidase2.041.021.82
metA Homoserine O-succinyltransferase3.843.453.98
metB Cystathionine gamma-synthase2.731.081.15
metK Methionine adenosyltransferase1.352.262.39
mhpA 3-(3-hydroxyphenyl)propionate hydroxylase3.801.281.23
mhpB 3-(2,3-dihydroxyphenyl)propionate dioxygenase4.241.461.39
mtlD Mannitol-1-phosphate 5-dehydrogenase1.671.242.03
nirD Nitrite reductase small subunit2.831.081.21
nuoC Bifunctional NADH:ubiquinone oxidoreductase subunit C/D2.001.661.55
nuoE NADH dehydrogenase subunit E2.021.541.50
nuoF NADH dehydrogenase I subunit F2.701.991.60
nuoG NADH dehydrogenase subunit G3.072.031.87
nuoI NADH dehydrogenase subunit I2.771.842.14
nuoM NADH dehydrogenase subunit M2.681.832.60
pabB Para-aminobenzoate synthase component I2.101.111.50
pdxA 4-hydroxy-l-threonine phosphate dehydrogenase, NAD-dependent2.321.221.14
phnP Carbon-phosphorus lyase complex accessory protein2.221.021.70
plsB Glycerol-3-phosphate acyltransferase2.561.131.05
purB Adenylosuccinate lyase2.091.11.27
purD Phosphoribosylglycinamide synthetase phosphoribosylamine-glycine ligase17.141.521.74
purE Phosphoribosylaminoimidazole carboxylase catalytic subunit4.381.091.61
purH Phosphoribosylaminoimidazolecarboxamide formyltransferase/IMP cyclohydrolase11.151.391.62
purK Phosphoribosylaminoimidazole carboxylase ATPase subunit4.051.141.77
purT Phosphoribosylglycinamide formyltransferase 26.781.281.65
puuA Glutamine synthetase2.031.221.51
puuD Gamma-glutamyl-gamma-aminobutyrate hydrolase2.441.471.50
puuR DNA-binding transcriptional repressor PuuR2.091.301.27
pyrB Aspartate carbamoyltransferase catalytic subunit2.011.371.03
pyrG CTP synthase2.021.321.29
pyrI Aspartate carbamoyltransferase regulatory subunit2.161.341.03
rbbA Fused ribosome-associated ATPase3.841.131.88
rbsK Ribokinase2.461.631.16
rbsR Transcriptional repressor RbsR2.051.231.17
rffM UDP-N-acetyl-d-mannosaminuronic acid transferase2.171.271.12
rffT 4-alpha-l-fucosyltransferase2.921.171.51
rlmB 23S rRNA (guanosine-2′-O-)-methyltransferase3.801.621.59
sdaB l-serine dehydratase1.572.511.12
sixA phosphohistidine phosphatase2.991.061.45
sodB Superoxide dismutase1.822.522.46
speC Ornithine decarboxylase2.061.291.46
spoU tRNA guanosine-2′-O-methyltransferase2.241.201.31
tauD Taurine dioxygenase1.052.371.19
tdcG l-serine dehydratase 12.561.241.24
thiD Phosphomethylpyrimidine kinase2.191.152.09
thiM Hydroxyethylthiazole kinase2.571.052.18
ubiX 3-octaprenyl-4-hydroxybenzoate carboxy-lyase3.861.121.49
uxaB Altronate oxidoreductase2.301.511.43
yfiD Autonomous glycyl radical cofactor GrcA1.264.261.77
Genes encoding proteins with diverse functions
cho Endonuclease of nucleotide excision repair2.001.161.66
csiE Stationary phase inducible protein CsiE2.131.021.41
dinB///dinP DNA polymerase IV2.701.401.92
dinG ATP-dependent DNA helicase4.511.461.88
dinI DNA damage-inducible protein I4.222.851.35
dnaE DNA polymerase III subunit alpha4.851.521.39
glpR DNA-binding transcriptional repressor GlpR2.751.331.94
hchA Chaperone protein HchA2.171.071.26
lldR DNA-binding transcriptional repressor LldR2.101.381.38
mazE Antitoxin MazE3.051.151.17
mazF Toxin ChpA3.061.071.27
mfd Transcription-repair coupling factor2.751.061.07
mutS Methyl-directed mismatch repair protein2.571.041.31
obgE GTPase ObgE2.021.481.46
parE DNA topoisomerase IV subunit B2.131.291.45
polB DNA polymerase II3.651.152.26
pspB Phage-shock protein B2.001.631.70
pspG Phage-shock protein G3.422.052.15
recA Recombinase A2.201.491.50
recB Exonuclease V (RecBCD complex), beta subunit3.451.031.04
recF Recombination protein F2.031.171.16
recG ATP-dependent DNA helicase2.971.061.24
recJ ssDNA exonuclease RecJ2.001.171.32
recN Recombination and repair protein3.861.871.68
recX Recombination regulator RecX4.983.092.34
rnhB Ribonuclease HII3.691.361.54
rnr Exoribonuclease R2.704.291.30
rseP Zinc metallopeptidase RseP2.021.041.06
sfa Cold-shock gene3.411.591.17
stpA DNA-binding protein, nucleoid-associated4.161.861.87
topA DNA topoisomerase I2.031.171.14
topB DNA topoisomerase III2.411.021.32
torD Chaperone protein TorD2.361.171.47
umuC DNA polymerase V, subunit UmuC9.173.823.06
umuD DNA polymerase V subunit UmuD9.004.083.32
uvrA Excinuclease ABC subunit A3.281.431.86
yaiL Nucleoprotein/polynucleotide-associated enzyme1.551.472.21
yebG DNA damage-inducible protein YebG2.701.721.45
Table 7. Genes showing mixed response to the overproduction of all three proteins
GeneGene product and/or function

↑csdA/wt

15 °C, 1 h

↑rnr/wt

15 °C, 1 h

↑pnp/wt

15 °C, 1 h

  1. Positive values indicate up-regulation and negative values indicate down-regulation. Changes more than 1.5-fold are considered statistically significant.

Genes involved in membrane synthesis/function
amtB Ammonium transporter6.15−1.211.22
appB Third cytochrome oxidase, subunit II1.17−1.292.33
appC Third cytochrome oxidase, subunit I1.04−1.312.40
artJ Arginine transporter subunit−4.87−1.041.13
blr Beta-lactam resistance membrane protein−2.721.02−1.24
chaA Calcium/sodium:proton antiporter−2.281.09−1.07
chaC Cation transport regulator2.23−1.261.44
cheA Chemotaxis protein CheA−1.312.70−1.57
cheB Chemotaxis-specific methylesterase−1.212.06−1.36
cheW Purine-binding chemotaxis protein−1.402.98−1.16
cheY Chemotaxis regulatory protein CheY1.012.51−1.39
cheZ Chemotaxis regulator CheZ1.172.05−1.05
clcA Chloride channel, voltage-gated2.57−1.101.17
cpxP Periplasmic repressor CpxP2.641.17−1.38
cusS Sensor kinase CusS2.20−1.021.24
cvpA Colicin V production protein5.72−1.001.75
degP Serine endoprotease1.58−1.84−4.63
evgS Hybrid sensory histidine kinase in two-component regulatory system with EvgA2.331.12−1.07
fecA Ferric citrate outer membrane transporter2.36−1.641.17
fecB Iron-dicitrate transporter subunit2.34−1.601.17
fecC Iron-dicitrate transporter subunit2.44−1.791.76
fecR Transmembrane signal transducer for ferric citrate transport2.08−1.761.10
fhuD Iron-hydroxamate transporter substrate-binding subunit2.17−1.251.47
fimB Tyrosine recombinase1.65−1.33−2.92
fimC FimC1.461.18−2.60
fimE Tyrosine recombinase−2.241.27−2.12
fimI FimI///fimbrial protein1.041.06−2.56
flgB Flagellar basal body rod protein FlgB−1.482.16−1.79
flgC Flagellar basal body rod protein FlgC−1.731.64−2.37
flgD Flagellar hook assembly protein−2.161.29−2.36
flgE Flagellar hook protein FlgE−2.141.45−2.05
flhC Transcriptional activator FlhC9.703.56−1.35
flhD Transcriptional activator FlhD5.322.87−1.43
fliA Flagellar biosynthesis sigma factor−1.082.37−1.25
fliC Flagellar filament structural protein (flagellin)1.892.93−1.02
ftsK DNA translocase FtsK2.77−1.031.07
ftsQ Cell division protein FtsQ2.43−1.051.16
glnL Nitrogen regulation protein NR(II)2.05−2.09−1.43
glpC sn-glycerol-3-phosphate dehydrogenase subunit C3.18−1.26−1.07
glpT sn-glycerol-3-phosphate transporter1.602.11−1.15
gspE General secretion pathway protein E2.18−1.101.10
gspI General secretory pathway component, cryptic2.58−1.151.46
gspO Bifunctional prepilin leader peptidase/methylase2.07−1.021.62
kefB Glutathione-regulated potassium-efflux system protein KefB−2.301.06−1.21
malF Maltose transporter subunit2.351.05−1.02
malM Maltose regulon periplasmic protein3.04−1.02−1.21
mdtD Multidrug efflux system protein3.08−1.021.81
mdtI Multidrug efflux system protein MdtI−1.831.152.00
mdtJ Multidrug efflux system protein MdtJ−2.011.051.80
mdtK Multidrug efflux protein−2.191.221.49
motA Flagellar motor protein MotA−1.102.72−1.46
motB Flagellar motor protein MotB−1.193.20−1.43
mreC Rod shape-determining protein MreC2.58−1.011.41
mreD Rod shape-determining protein MreD2.53−1.061.07
ompF Outer membrane protein F−2.381.201.41
ompT Outer membrane protease2.601.16−1.37
pbpC Penicillin-binding protein 1C3.05−1.101.28
phnE Membrane channel protein component of Pn transporter2.18−1.071.07
rbsB d-ribose transporter subunit RbsB4.101.89−1.35
rbsC Ribose ABC transporter permease protein5.031.95−1.11
rseB Periplasmic negative regulator of sigmaE2.19−1.43−1.64
rseC SoxR reducing system protein RseC2.55−1.26−1.69
sbp Sulfate transporter subunit−2.031.28−2.75
ssuB Alkanesulfonate transporter subunit−1.152.171.12
tap Methyl-accepting protein IV−1.382.30−1.33
tar Methyl-accepting chemotaxis protein II−1.323.14−2.34
tauA Taurine transporter substrate-binding subunit−1.113.69−1.25
tauB Taurine transporter ATP-binding subunit−1.103.44−1.28
thiQ Thiamin transporter subunit2.31−1.021.57
tsr Methyl-accepting chemotaxis protein I1.042.49−1.21
uhpT Hexose phosphate transporter3.31−1.021.42
uraA Uracil transporter−1.122.021.50
znuA High-affinity zinc transporter periplasmic component−2.051.18−1.19
Genes involved in cell metabolism
aceB Malate synthase1.15−1.08−2.05
aldA Aldehyde dehydrogenase A3.992.34−1.46
argA N-acetylglutamate synthase−1.32−1.032.00
argD Acetylornithine aminotransferase/succinyldiaminopimelate aminotransferase−2.25−1.071.42
argR Arginine repressor−2.01−1.081.04
astA Arginine succinyltransferase4.97−1.21−1.35
astB Succinylarginine dihydrolase7.98−1.11.08
astD Succinylglutamic semialdehyde dehydrogenase5.71−1.32−1.32
astE Succinylglutamate desuccinylase9.23−1.03−1.05
atoB Acetyl-CoA acetyltransferase2.02−1.061.21
cyaA Adenylate cyclase2.21−1.03−1.08
cysC Adenylylsulfate kinase−2.811.25−2.47
cysD Sulfate adenylyltransferase subunit 2−2.901.10−2.14
cysN Sulfate adenylyltransferase, subunit 1−2.201.11−1.76
entF Enterobactin synthase multienzyme complex component, ATP-dependent2.39−1.86−1.40
fruB Bifunctional fructose-specific PTS IIA/HPr protein1.793.47−1.24
fruK Fructose-1-phosphate kinase1.982.07−1.13
gcvP Glycine dehydrogenase///glycine dehydrogenase2.94−1.09−1.51
glcB Malate synthase G−2.861.98−3.57
glcD Glycolate oxidase subunit GlcD−1.932.41−3.80
glnA Glutamine synthetase2.14−1.75−1.22
glnG Nitrogen regulation protein NR(I)2.44−1.92−1.34
glnK Nitrogen regulatory protein P-II 222.4−1.181.39
glpK Glycerol kinase2.061.24−1.65
hcaR DNA-binding transcriptional regulator HcaR2.881.74−1.10
hemA Glutamyl-tRNA reductase2.11−1.091.60
hisI Fused phosphoribosyl-AMP cyclohydrolase/phosphoribosyl-ATP pyrophosphatase−2.18−1.11.03
ilvB Acetolactate synthase catalytic subunit2.281.62−1.11
ilvC Ketol-acid reductoisomerase−2.031.06−1.38
luxS S-ribosylhomocysteinase−2.231.091.06
malQ 4-alpha-glucanotransferase2.40−1.45−1.21
malT Transcriptional regulator MalT7.311.57−1.29
mazG Nucleoside triphosphate pyrophosphohydrolase2.08−1.121.07
menB Naphthoate synthase−2.09−1.091.35
metH Homocysteine-N5-methyltetrahydrofolate transmethylase, B12-dependent3.12−1.18−1.07
mglC Methyl-galactoside transporter subunit2.50−1.061.39
mobB Molybdopterin-guanine dinucleotide biosynthesis protein B2.02−1.091.14
nirB Nitrite reductase (NAD(P)H) subunit3.02−1.031.02
nrdE Ribonucleotide-diphosphate reductase subunit alpha1.11−2.74−3.36
nrdF Ribonucleotide-diphosphate reductase 2, beta subunit, ferritin-like1.54−4.02−5.62
ompA Outer membrane protein A−2.32−1.021.32
otsA Trehalose-6-phosphate synthase1.06−2.44−1.83
phnP Carbon-phosphorus lyase complex accessory protein−2.051.05−1.16
ppk Polyphosphate kinase2.00−1.31−1.36
ppsA Phosphoenolpyruvate synthase2.11−1.59−1.25
ppx Exopolyphosphatase2.03−1.39−1.42
purC Phosphoribosylaminoimidazole-succinocarboxamide synthase3.92−1.15−1.17
purF Amidophosphoribosyltransferase6.91−1.071.53
purL Phosphoribosylformyl-glycineamide synthetase6.31−1.141.21
purM Phosphoribosylaminoimidazole synthetase8.90−1.021.40
purN Phosphoribosylglycinamide formyltransferase4.41−1.211.13
pyrC Dihydroorotase3.52−1.021.13
recD Exonuclease V subunit alpha3.26−1.111.16
relA (p)ppGpp synthetase I/GTP pyrophosphokinase2.79−1.151.23
rpiB Ribose-5-phosphate isomerase B−1.011.122.23
rpmF 50S ribosomal protein L32−2.121.011.34
speB Agmatinase−2.051.081.45
speD S-adenosylmethionine decarboxylase−2.221.121.11
ssuD Alkanesulfonate monooxygenase−1.085.68−1.04
ssuE NAD(P)H-dependent FMN reductase−1.035.03−1.05
tnaA Tryptophanase3.992.25−1.10
torY Putative cytochrome C-type protein4.95−1.02−1.03
udp Uridine phosphorylase−1.282.02−1.53
wrbA TrpR binding protein WrbA1.152.34−1.07
Genes encoding proteins with diverse functions
cadC DNA-binding transcriptional activator−2.201.391.05
cdaR DNA-binding transcriptional activator1.95−2.27−2.18
deaD CsdA1.801.021.05
djlC Hsc56 co-chaperone of HscC2.68−1.011.37
hscC Putative DnaK protein2.09−1.061.42
ibpB Heat shock chaperone IbpB2.50−1.24−1.24
mhpR DNA-binding transcriptional activator MhpR2.19−1.041.16
pnp PNPase1.021.051.80
ptrA Protease III3.151.02−1.03
rseA Anti-RNA polymerase sigma factor SigE1.78−1.65−2.04
rpoE RNA polymerase sigma factor RpoE1.89−1.51−2.28
soxS DNA-binding transcriptional regulator SoxS3.162.13−1.23
xerC Site-specific tyrosine recombinase XerC2.03−1.061.08

As mentioned previously, Tables 1-4 present genes, which show up- or down-regulation in response to the deletion of either csdA, rnr or pnp. It may be speculated that low-temperature RNA metabolism of those genes, which are up-regulated by deletion of either of csdA, rnr or pnp may be influenced by these proteins. Thus, the absence of CsdA, RNase R or PNPase leads to higher levels of these target mRNAs. However, it is not clear why certain genes show down-regulation in the csdA null mutant strain or rnr or pnp deletion strains. It may be an indirect effect of the deletion of these genes. Alternative unknown mechanisms may also lead to such an observation under certain circumstances. Nonetheless, it is interesting that there are distinct sets of genes which are specifically down-regulated in response to the deletion of csdA, rnr or pnp. This may be of physiological significance with respect to the cold-shock response of cells.

Proteins encoded by genes most prominently differentially expressed in response to absence of CsdA/RNase R/PNPase are schematically presented in Fig. 3A,B. As described below, the analysis also shows that there are different sets of genes, which specifically respond to the deletion of either csdA, rnr or pnp.

Figure 3.

Schematic presentation of proteins encoded by genes most prominently, down-regulated (A) and up-regulated (B) in response to deletion of CsdA/RNase R/PNPase.

Genes that are more prominently changed
in the csdA null mutant strain

A number of genes and operons were down-regulated in response to the deletion of csdA including those involved in sugar or amino acid metabolism and transport (Fig. 3A). Interestingly, several of these showed up-regulation in cells overproducing CsdA suggesting that the response may be specific to the changes in the level of CsdA. malE involved in maltose transport that was down-regulated in all three deletions (Table 1), was previously (Phadtare & Inouye 2004) reported to be up-regulated in wild-type cold-shocked cells and down-regulated in cold-sensitive, quadruple csp deletion (ΔcspA ΔcspB ΔcspG ΔcspE) (Xia et al. 2001) cells indicating that it may be important for proper cold-shock response and cold acclimation of the cells. Other genes belonging to the mal operon such as, malG, malF malM and malT were also reported to be up-regulated in the wild-type cold-shocked cells and down-regulated in the quadruple csp deletion cells. Note that, in the present analysis, malE is down-regulated in all the three deletion strains, but is up-regulated only with CsdA and RNR overproduction (1.27 and 1.38, respectively), but not that of PNPase (-1.53). malG, malF, malM and malT were also up-regulated with the over-expression of CsdA (Table S2 in Supporting Information). In addition to the genes shown in Fig. 3A, copper homeostasis (cutC), a serine endoprotease (degP), dihydroxyacetone kinase (dhaK), cold-shock protein (sfa), (Table 4) are more prominently down-regulated in the csdA null mutant strain.

The possible target genes of CsdA may be those which show more significant up-regulation in the csdA null mutant strain and down-regulation with its over-expression. These were shortlisted from Tables 2, 3 and 5 and Tables S1 and S2 in Supporting Information and are presented in Fig. 3B, lower panel. Most of these genes showed similar response to RNase R, but not to PNPase. These genes are diverse and seem to be involved in processes ranging from cell metabolism, transport of metals to transcription regulation.

Genes that are specifically and significantly up-regulated in the Δpnp strain

Consistent with the fact that PNPase is the major ribonuclease active at low temperature several mRNAs showed up-regulation in its absence. Notably, the genes which encode proteins involved in chemotaxis (che operon, tap, tar and tsr) were previously reported to be important for cold-shock response of cells. Our previous analysis showed that, cheW and cheY genes were up-regulated in the wild-type cold-shocked cells and also transcription of cheW is presumably enhanced by Csp proteins (Phadtare et al. 2006). It was also reported that the tap, tar genes show prolonged induction in the wild-type strain at 15 °C (Phadtare & Inouye 2004). Thus, the present analysis shows that these mRNAs are furthermore increased in the Δpnp strain. These genes were down-regulated with the over-expression of PNPase. Note that these genes showed opposite response in that they were down-regulated in the csdA null mutant and Δrnr strains and (Table 3) and this effect was counteracted to some degree with their over-expression. A previous transcript analysis by Polissi et al. (2003) showed that PNPase degrades csp mRNAs at low temperature and these genes are up-regulated in the Δpnp strain. In the present analysis, several genes belonging to the cspA family show up-regulation in the Δpnp strain, but as the fold change was less than 2, these are only included in Table S1 in Supporting Information. These genes show down-regulation in the strain over-expressing PNPase (Table S2 in Supporting Information). Several other proteins encoded by genes that show significant up-regulation in the Δpnp strain, but not in the csdA null mutant or the Δrnr strains are shown in Fig. 3B, lower panel. These genes are diverse and seem to be involved mainly in transport or transcription regulation.

Although the DNA polymerase encoding genes such as dinB, umuC and umuD genes encoding recombination and repair proteins, and recN, recX and dinI genes encoding DNA damage-inducible protein show up-regulation in the Δpnp strain, this response is not specific as it is also observed in the cells overproducing PNPase. However, these gene show mostly down-regulation in the csdA null mutant and Δrnr strains (Tables 3 and 4) and significant up-regulation when CsdA or RNase R are overproduced; for example, both umuC and umuD increased 9-fold and recN, recX and dinI are increased ~fourfold in response to the CsdA over-expression (Table 6). Genes such as bglF (PTS enzyme), rhsC (RhsC protein) (Table 2), sulA (SOS cell division inhibitor), kch (potassium channel), ompN (outer membrane protein), hybD (hydrogenase endopeptidase) and yebG (DNA damage-inducible protein) also show up-regulation irrespective of deletion (Table 3) or over-expression (Tables S2 and Table 6) of PNPase.

Some of the other notable observations from the analysis of the deletion cells are, (i) several of the genes encoding flagellar proteins (flg) that are up-regulated in response to deletion of all three genes are up-regulated two to 12-fold upon cold shock and this induction was seen even at 3 h of cold shock (Phadtare & Inouye 2004). The flhD and fliA, the regulators of the flagellar operon (Liu & Matsumura 1994), are not significantly changed in csd null mutant and Δrnr strains, but are up-regulated in the Δpnp strain. Most of the flg and fli genes are down-regulated to some extent with the over-expression of CsdA, RNase R and PNPase. (ii) The fec operon involved in iron transport is not affected significantly (the change is below 1.5-fold) by the deletion of either csdA or pnp, but is increased in the Δrnr strain (Table 4). Interestingly, fecI and fecR, regulators of the fec operon (Van Hove et al. 1990), were not much changed in csdA null mutant and Δpnp, strains, but were up-regulated in the Δrnr strain. (iii) The sulfur metabolism genes were down-regulated in all three deletion strains. However, CysB, the known regulator of the cys regulon is not affected by these deletions.

The present analysis showed that there are distinct sets of genes, mRNAs of which showed elevated levels in the absence of CsdA and PNPase at low temperature. This may be the possible reason for cell growth inhibition caused by absence of these proteins at low temperature. This inhibition can only be relieved by the respective proteins. Thus, CsdA and PNPase cannot substitute for each other's function during cold-shock response and adaptation. Present analysis can provide explanation why RNase R can substitute for CsdA, albeit somewhat weakly, as several of the genes that show changes in response to the changes in the level of CsdA show similar response to RNase R deletion or over-expression. Additional effect on growth may be imposed by the fact that some of the genes that are down-regulated by either CsdA or PNPase deletion are reported to be essential for cell growth. Even though these are not completely absent in the csdA null mutant or Δpnp strain, a combination of decreased metabolism of large number of mRNAs and down-regulation of these essential genes may contribute toward cold sensitivity of individual deletion mutants. Interestingly, these genes are also mostly specific to either CsdA or PNPase. For example, cynS encoding cyanate hydratase is down-regulated almost fourfold in the Δpnp strain, but is moderately affected by the csdA deletion and is not affected by the rnr deletion (Table 1). Other essential genes such as cysW (sulfate/thiosulfate transporter subunit), nrdA and nrdB (ribonucleotide-diphosphate reductase), (Table 1), cheW (chemotaxis), hybD (hydrogenase 2 maturation endopeptidase), motA (flagellar motor protein) (Table 3) and cutC (copper homeostasis) (Table 4) are more significantly affected by the csdA deletion. Most of these are corrected by the respective overproduction of CsdA, RNase R or PNPase.

As mentioned above, Tables 5-7 present genes which respond to the overproduction of either of CsdA, RNase R or PNPase. These conditions were tested to furthermore narrow down the possible targets of each of the proteins, for example, if a gene shows up-regulation with deletion of PNPase and down-regulation with its overproduction, it is highly possible that PNPase may be involved the metabolism of its RNA at low temperature. It is also interesting to note genes which show concomitant increase in their levels with overproduction of CsdA, RNase R or PNPase.

Genes that are down-regulated in response to the overproduction of CsdA, RNase R or PNPase

Proteins encoded by genes which are more prominently down-regulated by the over-expression of CsdA as compared to that of RNase R or PNPase are shown in Fig. 4A. This inhibition is reversed in the csdA null mutant cells. Fig. 4A also shows genes which are specifically down-regulated by overproduction of RNase R or PNPase, the inhibition being reversed by the respective deletions. Ribonucleotide-diphosphate reductase (nrdE and nrdF) show significant down-regulation with the over-expression of RNase R or PNPase, but no change in response to the overproduction of CsdA (Table 7). The flagellar genes, which are up-regulated in the csdA null mutant, Δrnr and Δpnp strains are down-regulated in the CsdA and PNPase overproducing strains, but are not affected by RNase R overproduction (Table 7).

Figure 4.

Schematic presentation of proteins encoded by genes most prominently, down-regulated in response to overproduction of CsdA/RNase R/PNPase (A) and (A) and up-regulated in response to overproduction of CsdA/RNase R (B).

Genes that are up-regulated in response to the overproduction of CsdA, RNase R or PNPase

The proteins encoded by genes which are significantly and specifically up-regulated in response to the CsdA overproduction in comparison with RNase R and PNPase overproduction are shortlisted in Fig. 4B. Various other genes involved in cell metabolism such as bisZ, cynS, eutB, iap, rlmB and ubiX also belong to this category (Tables 6 and 7). Several of these genes were down-regulated in response in the csdA null mutant strain.

The gene hybD encoding an endopeptidase was up-regulated in both CsdA and RNase R overproduction, but not in the PNPase overproduction (Table 6). The proteins encoded by genes which are highly up-regulated specific to RNase R overproduction are also shown in Fig. 4B. In addition, genes involved in cell metabolism such as cdd, yfiD also belong to this category (Tables 6 and 7). This up-regulation was lost in the Δrnr strain. There are not many genes which are highly up-regulated specific to PNPase overproduction; two such genes are ada and alkB (Table 6). Genes such as dkgB and metA (cell metabolism), emrD (multidrug resistance), kch (potassium channel), umuC and umuD (DNA polymerase subunits) (Table 6) were highly up-regulated in response to the overproduction of CsdA, RNase R and PNPase. As mentioned above, umuC and umuD are down-regulated in the csdA null mutant and Δrnr strains, but show up-regulation in the Δpnp strain, so their response to the changes in the level of PNPase is not specific.

Conclusion

Previously, it was reported that CsdA does not exhibit strong specificity for RNA substrates in vitro, but it was suggested that it may have specificity for certain in vivo (Iost & Dreyfus 2006). Our previous data suggested that RNase R may have some preference for RNA substrates it can degrade (Awano et al. 2008). The observations that (i) RNase R can suppress the cold sensitivity of csdA null mutant strain, but not that of Δpnp strain and (ii) PNPase cannot suppress the cold sensitivity of the csdA null mutant strain strongly suggested that there must be distinct sets of target genes for each of these proteins which are relevant for their essential function at cold shock. A comparative DNA microarray analysis of the targets of CsdA, RNase R and PNPase at cold-shock condition was thus carried out. The main goal was to focus on genes which are up-regulated with deletion of either of the proteins and down-regulated with their over-expression. The present analysis showed that there are distinct sets of genes which show such responses to CsdA and PNPase at low temperature. There are a number of genes (for example, the che genes as discussed above) which are specifically up-regulated in the Δpnp strain and down-regulated in the strain over-expressing PNPase, consistent with the observation that PNPase is one of the major ribonucleases at low temperature. These are the target RNAs in the metabolism of which it may be playing an important role. Note that these genes do not show similar response to changes in the levels of CsdA or RNase R. This furthermore supports the notion that these are specific targets of PNPase at low temperature and that is why over-expression of either of these two proteins does not suppress the cold sensitivity of the Δpnp strain.

Present analysis showed that in the case of CsdA too, there exists a specific set of genes, which are up-regulated in the csdA null mutant strain and down-regulated with its over-expression. Several of these genes show similar response to RNase R deletion or over-expression, but not to PNPase. The genes responding to both CsdA and PNPase are diverse; however, the targets of PNPase seem to be mainly membrane, transport-related or transcription regulation genes, while those of CsdA are more diverse and are involved in the cell metabolism, transport and transcription regulation. PNPase is known to be part of the RNA degradosome at both optimal and low temperatures, whereas CsdA only at low temperature and RNase R is not part of this complex. Interestingly, PNP and CsdA are essential at only low temperature. Present analysis also showed that several of the genes that show changes in response to the changes in the level of CsdA show similar response to RNase R deletion or over-expression, but not to PNPase. This can explain why RNase R can compensate for CsdA function during cold-shock response and adaptation, but PNPase cannot. This suggests a possibility that in addition to being part of the degradosome at low temperature, CsdA has its own set of target mRNAs which need partial unwinding of their secondary structures by CsdA before they are degraded by endoribonucleases. These are not acted upon by PNP. Further work would involve detail bioinformatic analysis of these two sets of target genes to decipher if there are any common structural or sequence features within each group.

Experimental procedures

Bacterial strains and plasmids

E. coli wild-type strain JM83 [F-araΔ(lac-proAB) rpsL(strr)] (Yanisch-Perron et al. 1985) was used. Its csdA::kan, Δrnr and Δpnp strains are described previously (Yamanaka & Inouye 2001; Awano et al. 2007, 2008). The csdA::kan (designated as csdA null mutant) strain was constructed by insertion of kanamycin-resistant gene (1.3-kb fragment) into the middle of the coding region of the csdA gene, which completely inactivates it. The Δpnp strain was created by replacing the entire pnp gene with kanamycin cassette. The Δrnr strain was obtained from Keio collection, Japan and was constructed similarly as the Δpnp strain. The strains were grown in M9 medium supplemented with glucose (0.02–0.4%) and 0.4% Casamino Acids. Antibiotics such as ampicillin (50 μg/mL) (for cells harboring pINIII vectors) or chloramphenicol (50 μg/mL) (for the deletion strains) were supplemented as required. The isopropyl β-D thiogalactopyranoside (IPTG)-inducible pINIII plasmid and the pINIII-csdA, pINIII-rnr, and pINIII-pnp expression vectors were constructed by cloning the regions corresponding to the csdA, rnr and pnp, respectively, upon NdeI and BamHI and are described previously (Awano et al. 2007, 2008).

RNA isolation

E. coli cells grown overnight in M9 medium at 37 °C were diluted into fresh medium. The JM83 wild-type and the deletion strain cells were grown at 37 °C to exponential phase (OD600 of ~0.6), cold shocked at 15 °C for 1 h and were then harvested. To examine the effect of over-expression of respective proteins, the pINIII, pINIII-csdA, pINIII-rnr and pINIII-pnp plasmids were transformed into JM83 strain. The exponentially growing cells at an OD600 of 0.5 at 37 °C were then transferred to 15 °C and induced with 1 mm IPTG for 2 h. The cells were then harvested for RNA isolation. The total RNA was extracted by the hot phenol method described previously (Sarmientos et al. 1983). It was furthermore purified by RNeasy Minikit (Qiagen) and was then treated with DNase I followed by phenol-chloroform treatment and ethanol precipitation. It was quantified by measuring absorbance at 260 nm. The purity of RNA was confirmed by agarose gel electrophoresis.

DNA microarray analysis

The microarray analysis was carried out at the Center for Functional Genomics, University of Albany. This type of analysis has been successfully used by several laboratories (Bradley et al. 2007; Gorski et al. 2011; Jain et al. 2011). The RNAs were converted to sscDNA followed by fragmentation. The fragments are end-labeled with biotinylated GeneChip DNA labeling reagent and then hybridized to the arrays. GeneChip Ecoli genome 2.0 was used. The arrays are then washed, stained with streptavidin-phycoerythrin and scanned on a GeneChip scanner 30007G. The CEL files were imported into GenespringGX and quantile normalized upon PLIER. The signals were also baseline transformed to the median of all samples. The entities were filtered to exclude those with signal values less than 20th percentile across all conditions. This list was subjected to an anova (P < 0.05) with a Benjamini-Hochberg False discovery rate correction to identify statistical differences between all conditions versus control. The statistically significant genes were subjected to a 1.5-fold change filter to identify genes that were differentially expressed in each condition versus control. Differential expression was defined as equal to or greater than 1.5-fold change in expression in each set of control and test. We also used IntelliGene E. coli CHIP Version 2 (Takara Bio. Inc., Japan) (Phadtare & Inouye 2004; Phadtare et al. 2006) for analysis of Δpnp strain for additional confirmation of the data (not shown).

qRT-PCR

qRT-PCR was carried out in the Molecular Biology Core Facilities of the Center for Functional Genomics, University at Albany (Bradley et al. 2007). Genes to be tested for qRT-PCR were randomly selected from all the data tables (Tables 1-6) presented in the text. Reaction plates were processed on an Applied Biosystems 7900HT Sequence Detection System. The gene atpE identified by the microarray analysis was used an internal control. All reactions were carried out in triplicate. RE is to relative expression. The 7900HT software refers to RE as RQ (relative quantity). For the level of genes in the deletion cells, their respective level in wild-type is normalized to 1.0; for the level of genes in the cells over-expressing either CsdA, RNase R or PNPase, their respective level in wild-type cells carrying the vector alone is normalized to 1.0. To obtain the ± values, relative expression was calculated for the sum of the obtained RE value for a sample plus the ΔCt standard deviation. The RE± value obtained by this calculation was then subtracted from the RE value for this sample. Dissociation curve analysis verified the specificity of the primer pairs for each gene target, and there was no indication of primer-dimer formation in any reactions. No specific products were formed in the control reactions, which were carried out in the absence of templates. This was confirmed by both amplification plots and dissociation curve analysis.

Acknowledgements

This work was supported in part by NIH RO3 Grant 76900. I appreciate the critical reading of the manuscript by Dr Severinov. I would also like to acknowledge several excellent papers published on cold-shock response and cold-shock proteins from various groups, which could not be included here because of the lack of space.

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