1 Department of Biochemistry and Molecular Biology, University of Southern Denmark, Campusvej 55, 5230 Odense M, Denmark
2 Institut für Mikrobiologie und Molekularbiologie, Interdisziplinäres Forschungszentrum der Universität Giessen, Heinrich-Buff-Ring 26-32, 35392 Giessen, Germany
Correspondence
Lotte Søgaard-Andersen
sogaard{at}bmb.sdu.dk
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ABSTRACT |
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INTRODUCTION |
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Fruiting body morphogenesis involves temporally coordinated changes in gene expression in which genes are turned on at specific time points during development (Inouye et al., 1979; Kroos et al., 1986
). Moreover, for several genes that are turned on after 6 h, induction is tied to the spatial position of the cells, i.e. these genes are expressed in cells that aggregate and which eventually differentiate into spores, whereas they are not expressed in cells that differentiate to peripheral rods (Julien et al., 2000
). In contrast, genes activated prior to 6 h are expressed in all cells, including peripheral rods (Julien et al., 2000
). Three lines of evidence suggest that linked regulatory pathways coordinate morphogenesis and developmental gene expression (Søgaard-Andersen et al., 2003
). Firstly, most developmental mutants display defects in morphogenesis as well as in developmental gene expression (Dworkin, 1996
). Secondly, in several cases, developmentally regulated genes not only direct the expression of downstream developmental genes, but are also important for morphogenesis (e.g. Kroos et al., 1990
). Finally, the cell-position-specific expression of genes turned on after 6 h indicates that cells are able to detect their position during fruiting body formation and tailor their gene expression profile accordingly.
During fruiting body formation, cells interact with each other using at least five intercellular signals (A to E) (Shimkets, 1999). Analyses of developmental gene expression in signalling mutants suggest that the signalling systems are arranged in a time-based hierarchy and that they lie on the same developmental pathway (Cheng & Kaiser, 1989
; Downard & Toal, 1995
; Gill & Cull, 1986
; Kroos & Kaiser, 1987
; Kuspa et al., 1986
). Mutants deficient in any of the signalling systems are deficient in aggregation and sporulation, and display abnormal developmental gene expression. Examination of gene expression and fruiting body morphogenesis in signalling-deficient mutants has shown that the A and B signals become important early on in development, the D and E signals become important for development after 35 h, and the C signal becomes important for development after about 6 h.
Recent findings indicate that M. xanthus uses a wide array of different gene-regulatory proteins to direct developmental gene expression. These proteins include alternative sigma-factors (Apelian & Inouye, 1990, 1993
; Ueki & Inouye, 1998
, 2001
), DNA-binding response regulators (Ellehauge et al., 1998
; Ogawa et al., 1996
) including several NtrC-like activators (Caberoy et al., 2003
; Gronewold & Kaiser, 2001
, 2002
; Guo et al., 2000
; Hager et al., 2001
; Sun & Shi, 2001a
, b
), MrpC, a homologue of the E. coli CRP protein (Sun & Shi, 2001a
), and AsgB, which contains a C-terminal DNA-binding domain and a novel oligomerization and/or regulatory domain (Plamann et al., 1994
). Likewise, cis-acting regulatory sequences important for developmental gene expression are being revealed (Brandner & Kroos, 1998
; Fisseha et al., 1996
, 1999
; Gulati et al., 1995
; Hao et al., 2002
; Keseler & Kaiser, 1995
; Li et al., 1992
; Romeo & Zusman, 1991
; Ueki & Inouye, 2003
; Viswanathan & Kroos, 2003
; Yoder & Kroos, 2004
). Currently, the best-understood example of developmental gene expression at the molecular level involves the MrpC-dependent activation of the fruA gene, which encodes a DNA-binding response regulator that acts in the C-signal transduction pathway (Ellehauge et al., 1998
; Horiuchi et al., 2002
; Ogawa et al., 1996
; Søgaard-Andersen & Kaiser, 1996
). MrpC binds to the fruA promoter and activates fruA transcription after 36 h (Ueki & Inouye, 2003
). Interestingly, expression of mrpC is itself developmentally regulated (Sun & Shi, 2001b
). These observations suggest that developmental gene expression and the progression of the developmental programme in M. xanthus involve a complicated network of transcriptional regulators.
To further understand the mechanisms involved in developmental gene expression and the coupling between morphogenesis and developmental gene expression, we have analysed an M. xanthus mutant that displays delayed aggregation, reduced sporulation and abnormal developmental gene expression. Here, we report the identification of two co-transcribed genes that are important for normal aggregation, sporulation and developmental gene expression. The upstream gene hthA is likely to encode a novel DNA-binding protein. The predicted hthB product lacks homology to proteins in the databases. HthA is important for aggregation, whereas HthB, alone or in combination with HthA, is important for sporulation.
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METHODS |
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Bacterial strains and plasmids.
M. xanthus strains used in this work are listed in Table 1. SA1310, SA1625, SA1626, SA1627 and SA1628 were constructed by generalized transduction using Mx4 propagated on DK1622
0021, DK1622
2525, DK1622
4074, DK1622
4409 and DK1622
4695, respectively, to infect DK1622. SA1311, SA1312, SA1313, SA1315, SA1316, SA1317, SA1319, SA1323, SA1324 and SA1326 were constructed by generalized transduction using Mx4 propagated on SA1310 to infect DK1217, DK1300, SA1704, DK4293, DK4300, DK4368, DK4499, DK5279, DK9007 and DK11063, respectively. All strains constructed by generalized transduction were tested by Southern blot analyses (Sambrook et al., 1989
). SA1332, SA1333 and SA1335 were constructed by homologous integration of plasmid pMN304, pMN306 and pMN308, respectively, into the chromosome after electroporation (Kashefi & Hartzell, 1995
). Integration was verified by Southern blot analyses (Sambrook et al., 1989
). Plasmids used in this work are listed in Table 1
. pMN300 construction. A 6732 bp XhoI fragment containing miniTn5(tet)
0021 and flanking DNA sequences was cloned in the XhoI site in pBluescriptIISK (Stratagene). This fragment contains M. xanthus genomic DNA from 1896 to +1708 (all coordinates are relative to the start codon in hthA). pAAR106 construction. A 6732 bp XhoI fragment containing miniTn5(tet)
2525 and flanking DNA sequences was cloned in the XhoI site in pBluescriptIISK. This fragment contains M. xanthus genomic DNA from 1896 to +1708. pMN301 construction. From pMN300 a 1183 bp PstIXhoI fragment extending from +524 to +1708 was cloned in pBluescriptIISK. pMN302 construction. From pAAR106 a 2025 bp SacIPstI fragment extending from 1500 to +524 was cloned in the same sites in pMN301. pMN302 contains M. xanthus genomic DNA from 1500 to +1708. pMN304 construction. From pMN302 a 3205 bp SacIXhoI fragment extending from 1500 to +1708 was cloned in the same sites in pBGS18 (Spratt et al., 1986
). pMN306 construction. A 1691 AatIISalI fragment extending from position +978 to +2668 was generated by PCR using chromosomal DNA from DK1622 as a template and the primers hth7 and hth8 (primers used in this work are listed in Table 2
) followed by restriction with AatII and SalI. This fragment was cloned in the same sites in pMN302. From this pMN302 derivative a 4169 bp SacISalI fragment was cloned in the same sites in pBGS18 to generate pMN306. pMN306 contains M. xanthus genomic DNA from 1500 to +2668. pMN308 construction. A 404 bp blunt-end fragment extending from +1545 to +1948 was generated by PCR using chromosomal DNA from DK1622 as a template and the primers hth3 and hth4 (see Table 2
) followed by treatment with T4 DNA polymerase (New England Biolabs). This fragment was cloned in the SmaI site in pBGS18 to generate pMN308. All plasmids generated by PCR were verified by sequence analyses. Plasmids were propagated in TOP10 [F mcrA
(mrrhsdRMSmcrBC)
80lacZ
M15
lacX74 deoR recA1 araD139
(araleu)7679 galU galK rpsL endA1 nupG] (Invitrogen).
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RT-PCR.
To perform non-quantitative RT-PCR, RNA was isolated from vegetative cells. RNA was extracted by the hot-phenol method (Sambrook et al., 1989). The RNA was DNase I treated and re-extracted by phenol/chloroform extraction. Each of the primers hth2, hth4 and hth6 was used in a reverse transcription reaction with 100 ng total RNA using AMV RT (Finnzymes) according to the manufacturer's recommendations. Subsequently, the primer pairs hth1-2, hth3-4 and hth5-6 were used in the PCR reaction with Taq polymerase (Promega) according to the manufacturer's recommendations. To perform quantitative RT-PCR, cells were developed on CF agar for the indicated periods of time and harvested. RNA was extracted as described above. The RNA was reverse-transcribed using TaqMan Reverse Transcription Reagents with the supplied hexamers according to the protocol recommended by the supplier (Applied Biosystems). cDNA was purified using High Pure PCR Product Purification Kit (Roche). SYBR Green PCR Master Mix was added to cDNA from the reverse transcription of 100 ng RNA together with 500 nM of each of the two primers, hthF and hthR. The primers hybridize to the middle of hthA and give rise to a PCR product with a size of 65 bp (see Table 2
). The RT-PCR reaction was performed on an ABI PRISM 7700 Sequence Detection System (Applied Biosystems) using the standard set-up. Primers were designed using PrimerExpress as recommended by the ABI PRISM 7700 Sequence Detection System supplier. The level of hthAB transcript detected is expressed as relative units per ng total RNA.
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RESULTS |
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Two genetic systems control gliding motility in M. xanthus (Spormann, 1999). The social (S)-motility system controls gliding of groups of cells, and the adventurous (A)-motility system controls gliding of single cells. To test the effect of miniTn5(tet)
0021 on motility, the insertion was transduced into representative S (DK1300 sglG1) and A (DK1217 aglB1) backgrounds by Mx4-dependent generalized transduction. Strains that carry both an A mutation and an S mutation are non-swarming and grow as small, smooth-edged colonies (Hodgkin & Kaiser, 1979a
, b
). The miniTn5(tet)
0021 insertion did not generate a non-swarming phenotype when crossed into A or S mutant backgrounds (Fig. 2
). Thus, the miniTn5(tet)
0021 insertion does not interfere with either the A- or the S-motility system in vegetative cells, suggesting that the gene carrying the miniTn5(tet)
0021 insertion is not a constituent of either the A- or the S-motility system. Together, these data provide evidence that the developmental defects caused by the miniTn5(tet)
0021 insertion are not secondary to an effect on motility.
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hthA and hthB are co-transcribed
The overlap between the stop codon of hthA and the start codon of hthB suggested that these two genes are cotranscribed. To test this idea, non-quantitative RT-PCR analyses were carried out on total RNA isolated from vegetative wild-type cells using primer pairs specific for hthA and hthB, respectively, and a primer pair in which one primer hybridized to hthA and the second primer hybridized to hthB. All three primer pairs gave rise to a PCR product with the correct size in the RT-PCR reactions (Fig. 3b), thus suggesting that hthA and hthB constitute an operon.
Insertion of miniTn5(tet) into a gene is likely to inactivate it. Moreover, miniTn5(tet) may have polar effects on downstream genes. Thus, in the case of miniTn5(tet) 0021, the insertion may inactivate hthA and have a polar effect on hthB. To determine whether miniTn5(tet)
0021 has a polar effect on hthB transcription, RT-PCR analyses were carried out on total RNA isolated from vegetative SA1310 cells using the same three primer pairs as above. As shown in Fig. 3(b)
, no PCR products were detected in the RT-PCR reactions in these experiments. Thus, miniTn5(tet)
0021 insertion has a polar effect on hthB transcription.
Expression of the hthAB genes during development
To study the expression of the hthAB genes, quantitative RT-PCR analyses were performed on total RNA isolated from vegetative and starving DK1622 cells. The relative levels of hth mRNA decreased 500-fold between 0 and 6 h of starvation. After 12 h, the hth mRNA was not detectable in the RT-PCR analyses. As a control, the expression of the todK gene was also analysed by quantitative RT-PCR. As previously reported (Rasmussen & Søgaard-Andersen, 2003), expression of todK was observed to decrease 10-fold during the first 12 h of starvation. Thus, transcription of the hthAB genes decreases strongly in response to starvation.
Identification of the genes in the hth locus required for aggregation and sporulation
To determine whether the defects caused by the miniTn5(tet) 0021 insertion were due to loss of hthA and/or hthB function, genetic complementation experiments were performed. The plasmid pMN304 carries the wild-type hthA gene including 1500 bp upstream from the putative start codon of hthA (Fig. 3a
). The plasmid pMN306 carries the wild-type hthA and hthB genes including 1500 bp upstream from the putative start codon of hthA (Fig. 3a
). pMN304 and pMN306 were introduced by electroporation into SA1310, which carries the miniTn5(tet)
0021 insertion, to give strains SA1332 and SA1333, respectively. In both strains, the plasmid had integrated by homologous recombination upstream from the miniTn5(tet)
0021 insertion. SA1332 carries an intact hthA gene including the native promoter upstream from the integrated plasmid and the 3'-end of orfA, hthA : : miniTn5(tet)
0021 and hthB downstream from the integrated plasmid. SA1333 carries intact copies of hthA and hthB including the native promoter upstream from the integrated plasmid and the 3'-end of orfA, hthA : : miniTn5(tet)
0021 and hthB downstream from the integrated plasmid. SA1332 and SA1333 were assayed for development on CF starvation medium in parallel with DK1622 and SA1310. As shown in Fig. 1
and Table 3
, the aggregation defect caused by the miniTn5(tet)
0021 insertion was corrected by pMN304 in SA1332. However, pMN304 did not correct the sporulation defect caused by miniTn5(tet)
0021. On the other hand, both the aggregation defect and the sporulation defect caused by miniTn5(tet)
0021 were corrected by pMN306 in SA1333. These observations show that the developmental defects caused by miniTn5(tet)
0021 are due to a loss of both hthA and hthB function. Moreover, these data provide evidence that in a strain lacking both HthA and HthB, complementation with hthA restores the aggregation defect whereas hthB is required to restore the sporulation defect.
The results of the genetic complementation analyses suggested that HthB is only required for sporulation. To test this hypothesis, a strain that carries an hthB mutation was constructed. The plasmid pMN308 was constructed by cloning a 404 bp DNA fragment, which is located within hthB and extends from position +1545 to +1948 in hthB (Fig. 3a), in the plasmid pBGS18 that confers resistance to kanamycin. pMN308 was introduced into the wild-type DK1622 by electroporation to give strain SA1335. In SA1335, pMN308 has integrated by homologous recombination as a result of single crossover in the hthB gene. A single crossover yields kanamycin-resistant electroporants with two incomplete copies of hthB and is, therefore, likely to inactivate the hthB gene. To examine the possible developmental defects caused by the hthB : : pMN308 mutation, SA1335 was exposed to starvation on CF starvation medium in parallel with the wild-type strain DK1622 and SA1310, which carries miniTn5(tet)
0021. SA1335 had an aggregation phenotype similar to that of DK1622, and thus displayed no aggregation defects (Fig. 1
). However, the level of sporulation in SA1335 was reduced compared to that in DK1622 and similar to that of SA1310 after 72 h and 120 h. Taken together, these data show that loss of hthB function results in a sporulation defect.
Loss of hthA and hthB function does not interfere with synthesis of intercellular signals
To investigate whether the sporulation defect caused by miniTn5(tet) 0021 was cell-autonomous, cells of SA1310 were co-developed with an equal number of wild-type cells and the number of spores formed by the SA1310 cells was measured after 72 h of starvation. The sporulation defect in SA1310 remained unaffected by co-development with wild-type cells (Table 4
). Moreover, we found that SA1310 cells rescued sporulation in mutants lacking the A, B, C, D or E signal as efficiently as wild-type cells by extracellular complementation (Table 4
). Taken together, these observations suggest that the developmental defects caused by miniTn5(tet)
0021 are cell-autonomous and that miniTn5(tet)
0021 does not interfere with the synthesis of intercellular signals required for fruiting body morphogenesis.
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DISCUSSION |
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Analysis of the primary sequence of the C-terminal part of HthA showed that this part of HthA has similarity to the DNA-binding domain found in several DNA-binding proteins. Moreover, a detailed comparison of the C-terminal part of HthA to the DNA-binding domain in NarL revealed that the amino acid residues involved in maintaining the tertiary structure of the DNA-binding domain in NarL are conserved in HthA. Thus, at the structural level, HthA is predicted to contain a C-terminal helixturnhelix DNA-binding motif.
The activity of transcriptional regulators containing the DNA-binding domain found in HthA is modulated in distinct ways: the activity of the FixJ-like DNA-binding response regulators is modulated by phosphorylation of a conserved Asp residue in the N-terminal receiver part of the proteins (Parkinson & Kofoid, 1992); binding of the cognate AHL autoinducer to the N-terminal domain modulates the transcriptional activity of the LuxR-like transcriptional regulators (Fuqua et al., 2001
); the GerE protein only consists of the four
-helix bundle DNA-binding domain and the activity is not modulated by either covalent modification or ligand binding (Ducros et al., 2001
); finally, the MalT protein in E. coli is activated by binding of ATP and maltotriose (Raibaud & Richet, 1987
; Richet & Raibaud, 1989
). Intriguingly, the N-terminal part of HthA does not exhibit significant similarity to any proteins in the databases. This lack of homology raises the possibility that HthA contains a novel domain involved in modulating transcriptional activity. Alternatively, this part of HthA could be involved in oligomerization, as has been found in TraR (Zhang et al., 2002
). In conclusion, the data support the notion that HthA contains a C-terminal DNA-binding domain and may act as a transcriptional regulator. This hypothesis is supported by the observation that loss of HthA and HthB function alters the gene expression profile in vegetative as well as in starving cells (see below).
The primary sequence of the HthB protein does not share significant homology to proteins in the databases. Therefore, the mode of action of the HthB protein in the sporulation process remains unknown. Likewise, our data do not allow us to conclude whether HthB functions independently of HthA during sporulation. In principle, HthA and HthB could act independently during fruiting body formation, with HthA being important for aggregation and HthB being important for sporulation. HthA and HthB could also interact directly or indirectly to promote sporulation. Given that the activity of some of the transcriptional regulators containing the DNA-binding domain found in HthA is regulated by ligand binding, an interesting possibility for an indirect interaction between HthA and HthB could be that HthB possesses enzymic activity and is involved in the synthesis of an HthA ligand, which is required for full activity of HthA. Alternatively, HthB may be required for full expression of hthA. Further experiments are needed to discriminate between these possibilities.
RT-PCR analyses provided evidence that the hthA and hthB genes are co-transcribed. No ORFs were identified in the 710 bp between the stop codon in orfA and the start codon of hthA (Fig. 3), suggesting that hthA and hthB may constitute a two-gene operon. hthA and hthB are expressed in vegetative cells. Accumulation of the hthAB mRNA decreases approximately 500-fold within the first 6 h of starvation, and after 12 h, the hthAB mRNA is no longer detectable. Assuming that the intracellular concentration of the HthA and HthB proteins follows the detection profile of hthAB mRNA, then HthA and HthB are present in vegetative cells and the cellular concentration of the two proteins decreases in response to starvation. Consistent with the notion that HthA and HthB are present in vegetative cells, an hthAhthB mutation results in increased transcription of the transcriptional sdeKlacZ fusion (Tn5lac
4408) in vegetative cells (see below).
Loss of HthA and HthB function has pleiotropic effects on gene expression. Loss of HthA and HthB function results in increased expression of a transcriptional sdeKlacZ fusion (Tn5lac 4408) in non-starving cells and during development. SdeK encodes a histidine protein kinase important for fruiting body formation (Garza et al., 1998
; Pollack & Singer, 2001
) and, normally, expression of sdeK is activated early during development in a stringent-response-dependent manner from a putative sigma-54-dependent promoter (Garza et al., 1998
; Singer & Kaiser, 1995
). The effect of loss of HthA and HthB function on sdeK expression provides evidence that HthA and/or HthB directly or indirectly inhibit transcription of sdeK in vegetative cells and in developing cells. Loss of HthA and HthB function also results in decreased levels of expression of transcriptional fruAlacZ (Tn5lac
7540) and devRlacZ fusions (Tn5lac
4414) during development. fruA is normally expressed after 36 h of starvation and encodes a DNA-binding response regulator, which is a key component in the C-signal transduction pathway (Ellehauge et al., 1998
; Horiuchi et al., 2002
; Ogawa et al., 1996
; Søgaard-Andersen & Kaiser, 1996
). devR is normally expressed after 69 h of starvation and encodes a protein of unknown function, which is important for sporulation (Thöny-Meyer & Kaiser, 1993
). These data argue that HthA and/or HthB directly or indirectly stimulate transcription of fruA and devR. Loss of SdeK function does not change expression of fruA (A. Aa. Rasmussen, unpublished), suggesting that the decreased transcription of fruA in the hthAhthB mutant is not caused by the increased levels of SdeK. On the other hand, both SdeK and FruA stimulate transcription of the devRlacZ fusion (Tn5lac
4414) (Ellehauge et al., 1998
; Kroos et al., 1990
) and Tn5lac
4403 (E. Ellehauge, unpublished; Pollack & Singer, 2001
). Therefore, the expression profile of the devRlacZ fusion (Tn5lac
4414) in the hthAhthB mutant could be explained by the lack of FruA protein being dominant over the increased level of SdeK. The observation that the expression profile of Tn5lac
4403 in the hthAhthB mutant is similar to the expression profile in wild-type cells could be explained by the increase in SdeK and decrease in FruA levels balancing each other.
How does loss of HthA and HthB function result in developmental defects? Complete loss of FruA function results in aggregation and sporulation defects (Ellehauge et al., 1998; Ogawa et al., 1996
). Likewise, loss of devR function results in a sporulation defect (Kroos et al., 1990
). We speculate that the decreased expression of fruA and devR contributes to the aggregation and sporulation defects in the hthAB mutant. Based on the strong decrease in transcription of the hthAB genes in response to starvation it is tempting to speculate that HthA and/or HthB may have their primary function in vegetative cells and only directly regulate gene expression in vegetative cells. In this scenario, proteins encoded by those genes that are regulated by HthA and/or HthB in vegetative cells would, in turn, be involved in the expression of genes important for aggregation and sporulation. Specifically, genes expressed in an HthA-dependent manner in vegetative cells would direct the expression of genes important for aggregation. Similarly, genes expressed in an HthB-dependent manner in vegetative cells would direct the expression of genes important for sporulation. In ongoing experiments we are addressing the identification of genes that are directly regulated by HthA and/or HthB.
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ACKNOWLEDGEMENTS |
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Received 8 March 2004;
revised 26 April 2004;
accepted 28 April 2004.
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