Deletion of HVO_1527, 1528 or 1529 does not compromise H. volcanii survival
As a first step in assessing the putative involvement of HVO_1527, 1528 and 1529 in H. volcanii protein glycosylation, each gene was deleted according to the protocol developed by Allers et al. (2004) and successfully used by numerous laboratories for the study of a variety of genes (cf. Soppa et al., 2008). In this approach, the sequence under consideration is replaced by the tryptophan synthase-encoding H. volcanii trpA gene (HVO_0789), introduced into the genome of the uracil and tryptophan auxotrophic H. volcanii strain WR536 by the pyrE-containing plasmid pTA131 and plating onto casamino acids lacking uracil and tryptophan.
PCR amplification was performed to follow genomic integration of the introduced plasmids as well as the subsequent expulsion of the plasmid together with HVO_1527, 1528 or 1529. However, given the comparable sizes of these genes and the trpA sequence (732, 888, 942 and 834 nucleotides, respectively), each gene was followed by dual PCR amplifications using forward primers raised against internal sequences within HVO_1527, 1528, 1529 or trpA and a reverse primer directed against a sequence within the flanking region downstream to HVO_1527, 1528 or 1529, as appropriate. As revealed in Fig. 1B (left panels), whereas those primer pairs directed against internal and downstream flanking regions of HVO_1527, 1528 or 1529 yielded PCR amplification products in the background strain (right pair of lanes in each panel; 1467, 1422 and 1251 bp, respectively), only those primer pairs directed against an internal sequence of trpA and the flanking regions downstream to HVO_1527, 1528 or 1529 yielded PCR amplification products in the deletion strain (left pair of lanes in each panel; 1353, 1368 and 1359 bp, respectively). These results thus point to respective replacement of HVO_1527, 1528 and 1529 by trpA. Deletion of each gene was further confirmed when PCR amplification was performed using genomic DNA from the deletion strains as template and primers directed against the HVO_1527, 1528 or 1529 coding regions (732, 888 and 942 bp, respectively; Fig. 1B, right panels).
Figure 1. HVO_1527, 1528 and 1529 are not essential for H. volcanii survival. A. Schematic representation showing the orientations of HVO_1527, 1528, 1529 and 1530 as well as proven (HVO_1530) annotations. B. Left panels: PCR amplification was performed using a forward primer directed at the HVO_1527, 1528 or 1529 3′ flanking regions and a reverse primer directed at a sequence within the HVO_1527, 1528 or 1529 coding regions (primer pair a) or a sequence within the trpA sequence (primer pair b), together with genomic DNA from cells of the WR536 background strain (bkgnd) or from cells that had replaced the HVO_1527, 1528 or 1529 gene (deletion; top, middle and bottom panels, respectively), as template. Right panel: PCR amplification was performed using primers directed against the HVO_1527, 1528 or 1529 coding regions, together with genomic DNA from cells of the WR536 background strain (bkgnd) or the HVO_1527, 1528 or 1529-deleted strains (deletion; top, middle and bottom panels, respectively). C. RT-PCR was performed using primers directed at HVO_1527 (top row of panels), HVO_1528 (middle row of panels) or HVO_1529 (bottom row of panels) and cDNA (left lane of each panel), RNA (middle lane of each panel) from HVO_1527, 1528 or 1529-deleted strains (left, middle and right columns of panels, respectively) as template. In the right lane of each panel, no nucleic acid template was added to the reaction (blank). The identities of PCR products were confirmed by sequencing.
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The absence of HVO_1527, 1528 or 1529 in the respective deletion strains was next ascertained at the RNA level by RT-PCR, performed as described previously (Abu-Qarn and Eichler, 2006). In these experiments, RNA or cDNA generated from the RNA of each deletion strain or no nucleic acids (blank) served as template for PCR amplifications, together with primers directed against the coding region of HVO_1527, 1528 or 1529. As reflected in Fig. 1C, no PCR products were obtained when cDNA from any of the deletion strains served as template in reactions involving primers directed against the deleted gene in question. By contrast, PCR products were readily obtained when the same reactions were repeated using primers directed against either of the other two sequences. For example, when PCR amplification was performed using cDNA obtained from cells deleted of HVO_1527 (Fig. 1C, left panels, left lanes), PCR products were obtained when primers to the HVO_1528 or 1529 coding regions were employed (middle and bottom panels, respectively) but not when primers to the HVO_1527 coding region were included in the reaction (top panel). Similarly, no PCR products appeared when RNA served as template (middle lanes of each panel) or when no nucleic acids were present (blank, right lanes of each panel). These results thus reflect the deletion of HVO_1527, 1528 and 1529 at the RNA level and, moreover, reveal that the absence of HVO_1527, 1528 or 1529 does not compromise the transcription of the other two genes.
H. volcanii cells lacking HVO_1527, 1528 or 1529 present a modified S-layer
Having determined that HVO_1527, 1528 and 1529 are not essential for H. volcanii viability, the participation of the gene products in protein glycosylation was considered by examining the S-layer glycoprotein, a well-characterized archaeal reporter of this post-translational modification (Sumper et al., 1990; Mengele and Sumper, 1992, Eichler, 2000), in cells deleted of each gene. As reflected in Fig. 2A, when the S-layer glycoprotein from the H. volcanii WR536 strain background and the same strain lacking HVO_1527, 1528 or 1529 were compared by SDS-PAGE and Coomassie staining, the faster migration of the protein from the mutant cells was evident. To confirm that such enhanced migration of the S-layer glycoprotein on SDS-PAGE was due to the absence of the individual genes and not the outcome of a general perturbation of the genome in the region of HVO_1527, 1528 and 1529, each deletion strain was transformed to express a plasmid-based copy of the absent gene, engineered to include a cellulose-binding domain (CBD) tag, with expression being confirmed by immunoblot using anti-CBD antibodies (Fig. 2B). In the case of HVO_1527 and 1529, such complementation restored the original SDS-PAGE behaviour of the S-layer glycoprotein (Fig. 2C). The failure of plasmid-encoded CBD-HVO_1528 to restore S-layer glycoprotein migration in SDS-PAGE to that of the native protein may be due to several causes, including steric interference by the fused CBD tag, introduced for purposes of detection. Nonetheless, the observation that HVO_1527 and HVO_1529 mRNA is detected in the HVO_1528 deletion strain (Fig. 1C) argues that effects resulting from the absence of HVO_1528 (such as modified S-layer glycoprotein apparent molecular weight) are due to the missing gene product rather than arising in a non-specific, unrelated manner because of genome disruption.
Figure 2. The absence of HVO_1527, 1528 or 1529 affects S-layer glycoprotein migration on SDS-PAGE. A. Equivalent aliquots of H. volcanii WR356 cells (bkgnd), or the same cells lacking HVO_1527 (ΔHVO_1527; top panel), HVO_1528 (ΔHVO_1528; middle panel) or HVO_1529 (ΔHVO_1529; bottom panel) were separated by 5% SDS-PAGE and Coomassie blue-stained. The position of the S-layer glycoprotein is shown. B. The expression of CBD-tagged HVO_1527 (top panel), HVO_1528 (middle panel) and HVO_1529 (top panel) in the complemented deletion strains, as confirmed by immunoblot after separation on 15% SDS-PAGE using anti-CBD antibodies. C. Equivalent aliquots of H. volcanii WR356 cells lacking HVO_1527 (ΔHVO_1527), HVO_1529 (ΔHVO_1529) or cells of the deletion strain transformed with a plasmid encoding a CBD-tagged version of the deleted gene (ΔHVO_1527/CBD-HVO_1527;ΔHVO_1529/CBD-HVO_1529) were separated by 5% SDS-PAGE and Coomassie blue-stained. The position of the S-layer glycoprotein is shown.
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To determine whether the enhanced migration of the S-layer glycoprotein in the HVO_1527-, 1528- or 1529-lacking cells was indicative of modifications that affected the integrity of the S-layer surrounding H. volcanii, thought to be composed solely of the S-layer glycoprotein (Sumper et al., 1990), WR536 background cells, as well as cells of the same strain deleted of HVO_1527, 1528 or 1529, were challenged with proteinase K for up to 3 h. The proportion of non-digested S-layer glycoprotein remaining at increasing intervals from the onset of proteolysis was then considered. Such analysis revealed the S-layer glycoprotein in the background strain (as well as in cells deleted of HVO_A0586, a seemingly unrelated putative nucleoside diphosphate sugar pyrophosphorylase) to be less susceptible to proteolytic digestion than its counterparts in the HVO_1527-, 1528- or 1529-deleted strains (Fig. 3). Moreover, complementation of HVO_1527- or HVO_1529-deleted cells to express a CBD-tagged version of the encoded protein restored the protease resistance of the S-layer to that of the background strain. It thus appears that the source of the enhanced HVO_1527-, 1528- and 1529-deleted strain-derived S-layer glycoprotein SDS-PAGE migration also compromises the proper assembly of the protein shell surrounding H. volcanii cells in these strains.
Figure 3. The S-layer surrounding H. volcanii cells is protease-sensitive in cells lacking HVO_1527, 1528 or 1529. Background strain WR536 (top panel), and HVO_1527-, 1528- or 1529-lacking cells of the same strain (second, third and fourth panels, respectively) were challenged with 1 mg ml−1 proteinase K at 42°C. In the fifth and sixth panels, cells lacking HVO_1527 or HVO_1529, respectively, transformed to express a CBD-tagged version of the deleted gene, were similarly challenged. Aliquots were removed immediately prior to incubation with proteinase K and at 15–30 min intervals following addition of the protease for up to 3 h and examined by 7.5% SDS-PAGE. In a control experiment, H. volcanii cells deleted of a seemingly non-related gene (HVO_A0586) were similarly challenged (bottom panel).
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HVO_1527, 1528 and 1529 participate in the assembly of the pentasaccharide decorating the H. volcanii S-layer glycoprotein
As homology-based predictions assign the deduced products of HVO_1527, 1528 and 1529 roles in N-glycosylation, experiments were next performed to directly test these predictions. Indeed, defective N-glycosylation could explain the observations described above. Accordingly, SDS-PAGE gel pieces containing the S-layer glycoprotein from the H. volcanii background strain, as well as from cells lacking HVO_1527, 1528 or 1529, were subjected to in-gel tryptic digestion. The obtained peptides were separated by liquid chromatography and MS/MS was employed to reveal peptide sequences. The six peptides generated in this manner included the N-terminal 1ERGNLDADSESFNK14 peptide (1581 m/z), encompassing the glycosylated Asn-13 residue (Sumper et al., 1990; Abu-Qarn et al., 2007). Previous MALDI TOF mass mapping of the nanoLC-purified tryptic digest, complemented by MS/MS analyses using MALDI TOF/TOF and electrospray Q-TOF instrumentation, had shown this S-layer glycoprotein-derived peptide to be modified by a novel pentasaccharide (Abu-Qarn et al., 2007). In agreement with this earlier study, the Asn-13-containing peptide isolated from the WR536 background strain was now shown to be decorated by the same pentasaccharide moiety (m/z 2447), composed of a hexose (162 Da), followed by two 176 Da residues (hexuronic acids), one 190 Da residue (likely either dimethylated hexose or the methyl ester of hexuronic acid) and an additional hexose residue at the end of the glycan chain. In addition, the same peptide modified by precursor mono- (m/z 1743.7), di- (m/z 1919.7), tri- (m/z 2095.8) and tetrasaccharides (m/z 2285.5) were also observed (Fig. 4, top left panel, bkgnd).
Figure 4. The products of H. volcanii HVO_1527, 1528 and 1529 are involved in the biogenesis of the pentasaccharide decorating S-layer glycoprotein Asn residues. The MALDI-TOF spectra of the Asn-13-containing tryptic peptide derived from the S-layer glycoprotein of the WR536 background cells (upper left panel) and cells from the HVO_1527- [ΔHVO_1527 (aglF); upper right panel], HVO_1528- [ΔHVO_1528 (aglI); lower left panel] or HVO_1529- [ΔHVO_1529 (aglG); lower right panel] deleted strains are shown. The components of the peptide-associated glycan are shown as an inset in the upper left panel, while the sugar subunits decorating the peptide peaks detected are indicated on the MALDI-TOF spectra.
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When the same S-layer glycoprotein peptide was derived from H. volcanii cells deleted of HVO_1527, a very different profile was obtained. In this case, the monosaccharide-bearing species as well as a lesser amount of the disaccharide-bearing peptide were observed [Fig. 4, top right panel, ΔHVO_1527 (aglF)]. In the case of cells deleted of HVO_1528, an identical pattern was obtained [Fig. 4, bottom left panel, ΔHVO_1528 (aglI)]. Due to the weak nature of the signal at m/z 1919.7 in the ΔHVO_1527 (aglF) and ΔHVO_1528 (aglI) profiles, relative to the m/z 1743.8 signal, two further analyses were performed (data not shown). Standard deviations of ±2.2 and ±0.9 were obtained for the relative intensities between these signals in the ΔHVO_1527 (aglF) and ΔHVO_1528 (aglI) samples, respectively. When the N-terminal S-layer glycoprotein tryptic peptide from cells lacking HVO_1529 was examined as above, only the monosaccharide-decorated species was detected [Fig. 4, bottom right panel, ΔHVO_1529 (aglG)]. The products of HVO_1527 and 1528 are therefore involved in the addition of the distal hexuronic acid of the pentasaccharide decorating Asn-13, while the product of HVO_1529 is involved in addition of the proximal hexuronic acid of the same oligosaccharide.
Thus, given the involvement of HVO_1527, 1528 and 1529 in H. volcanii S-layer glycoprotein N-glycosylation, as revealed by analysis of SDS-PAGE migration, susceptibility to proteolysis and mass spectrometry of this reporter, HVO_1527, 1528 and 1529 are now renamed aglF, aglI and aglG, according to the nomenclature proposed by Chaban et al. (2006) for genes involved in archaeal N-glycosylation.
The transcription of aglB, aglF, aglG and aglI is regulated in a co-ordinated manner
Given their physical proximity in the genome, as well as the common involvement of their products in N-glycosylation, efforts next focused on whether the transcription of aglB, aglF, aglG and aglI is co-ordinated. Towards this end, the transcription profile of these genes was initially considered by investigating their upstream regions for the presence of promoters.
As aglF and aglI lie adjacent to each other on the H. volcanii genome, assume the same orientation and are apparently separated by only 51 nucleotides, the possibility that the two genes are co-transcribed was tested. Accordingly, RT-PCR was performed using cDNA derived from RNA extracted from cells grown to mid-exponential phase, together with a forward primer directed against the 5′ end of aglF and a reverse primer directed against the 3′ end of aglI. As reflected in Fig. 5A, a single PCR product was obtained, confirmed by sequencing to contain both aglF and aglI. To further demonstrate that the transcription of aglF and aglI is under the control of a common promoter, the DNA sequence separating HVO_1526 and 1527, i.e. that region lying upstream of the predicted start site of aglF, was introduced into plasmid pJAM1020 (Reuter and Maupin-Furlow, 2004), encoding for GFP, in place of the Halobacterium cutirubrum rRNA P2 promoter originally present in the plasmid. Preliminary control experiments confirmed that in the absence of the native promoter, no GFP expression could be detected (Fig. 5B). When the modified plasmid containing the 180 bp region upstream of the predicted aglF start site in place of the original promoter was used to transform H. volcanii cells, the expression of GFP could be clearly seen. By contrast, far less GFP expression was achieved when the original plasmid pJAM1020 promoter region was replaced with the 51 bp sequence separating aglF and aglI. Thus, although the region upstream of aglI is capable of directing protein expression to a limited extent, the augmented level of protein expression directed by the stronger aglF promoter offers additional support for the concept that aglF and aglI, the products of which jointly participate in addition of a hexuronic subunit to the S-layer glycoprotein pentasaccharide, are co-transcribed under the control of a single promoter.
Figure 5. Functional characterization of the promoter regions of H. volcanii aglB, aglF, algG and aglI. A. RT-PCR reveals the co-transcription of aglF and aglI. PCR amplification was performed using a forward primer directed against the start of the coding region of aglF and a reverse primer against the end of the coding region of aglI together with cDNA (lane 1), RNA (lane 2) or DNA (lane 3) from H. volcanii strain WR536 background cells as template, or no nucleic acid template (lane 4). B. Upper panels: H. volcanii strain WR536 cells were transformed to express GFP, as directed by plasmid pJAM-1020 in which the promoter region had been removed, in which the native promoter was present, or when the region upstream to aglF or aglI replaced the native promoter of the plasmid. Lower panel: The 118 bp region separating aglG and aglB served as promoter (aglB lane). In lane aglG, the same region, this time in the reverse orientation, served as the promoter in plasmid pJAM-1020. GFP expression was visualized by immunoblotting using anti-GFP antibodies.
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Unlike aglF and aglI, which assume the same orientation in the H. volcanii genome, aglG and aglB are oriented in opposite directions, with the direction of aglG being inverted. To determine whether the 118 bp DNA sequence separating the predicted start sites of aglG and aglB drives the transcription of either or both genes, the original plasmid pJAM1020 promoter was replaced by the 118 bp region, introduced in either orientation. As also reflected in Fig. 5B, the transcription of GFP was driven by this H. volcanii sequence, regardless of its orientation. The level of GFP detected was, however, somewhat higher when the 118 bp region was inserted in the forward direction, pointing to the promoter being stronger in driving the transcription of aglB than of algG.
While aglB, aglF, aglG and aglI do not form an operon, given the differential orientation of aglG, the possibility remains that their transcription is somehow linked. To begin testing this hypothesis, the relative levels of aglB, aglF, aglG and aglI transcription were qualitatively assessed by comparing the levels of GFP in cells transformed to transcribe the encoding gene under the control of the aglB, aglF, aglI or aglG promoters, grown to mid-exponential phase. As reflected in Fig. 5, GFP expression driven through the aglB promoter exceeded that expression directed through the aglF, aglI or aglG promoters. To quantify these observations, real-time RT-PCR was performed to assess the relative amounts of AglB, AglF, AglG and AglI mRNA in H. volcanii cells grown to mid-exponential phase in complete medium, using primer pairs that bind with equivalent efficiencies (as determined in preliminary experiments involving the drawing of standard curves describing primer pair binding to serial dilutions of known quantities of cDNA). Based on the results of three experiments, each involving triplicate samples, it could be concluded that H. volcanii cells grown to mid-exponential phase in complete medium contain threefold less AglF mRNA, and fivefold less AglG and AglI mRNA than AglB mRNA (with all differences being significant to P < 0.01) (Fig. 6A). At present, it is not clear why different levels of AglF and AglI mRNA were detected, if the two genes are co-transcribed. Thus, cells grown to mid-exponential phase contain different amounts of aglB, aglF, aglG and aglI mRNA.
Figure 6. The transcription of aglB, aglF, algG and aglI is co-ordinated. A. Real-time RT-PCR was employed to assess the relative amounts of aglB, aglF, algG and aglI mRNA in H. volcanii strain WR536 cells grown to mid-exponential phase in rich medium. Values shown represent the average of three experiments ± standard deviation, expressed relative to the level of aglB mRNA, taken as 1. Differences from the level of aglB RNA marked with the double asterisk are statistically significant to P < 0.01, as determined by Student's t-test. B. Real-time RT-PCR was employed to assess the fold increase in aglB, aglF, algG and aglI mRNA in H. volcanii strain WR536 cells grown to stationary phase (stat.), subjected to heat shock, or raised in low or high salt-containing medium, relative to those levels detected in cells grown to mid-exponential phase in rich medium. The values shown represent the average of 3–5 experiments. Bars marked with the double asterisk are statistically distinct to a significance of P < 0.01, while those marked with single asterisks are statistically distinct to a significance of P < 0.05, as determined by Student's t-test.
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To determine whether changes in the transcription profile of aglB, aglF, aglG and aglI take place in a co-ordinated manner, real-time PCR was employed to quantify AglB, AglF, AglG and AglI mRNA levels in cells grown to stationary phase, challenged with heat shock (i.e. 65°C for 45 min) or raised to mid-exponential phase in low salt- or high salt-containing medium (i.e. 1.75 or 4.8 M NaCl, respectively). In these experiments, 16S rRNA was considered as a housekeeping marker to allow for normalization of mRNA levels from cells in each growth condition. The AglB, AglF, AglG and AglI mRNA levels measured in these various growth conditions were, in turn, expressed in terms of fold increase relative to those values obtained from cells grown to mid-exponential phase in complete medium. Initially, AglB, AglF, AglG and AglI mRNA levels in cells either grown to stationary phase or exposed to heat shock conditions are considered. In cells grown to stationary phase, AglB, AglF, AglG and AglI mRNA levels were substantially reduced, relative to those levels realized during mid-exponential growth (Fig. 6B). AglB, AglF, AglG and AglI mRNA levels were also depressed upon transfer to heat shock conditions, relative to the situation realized in cells grown to mid-exponential phase. In both growth conditions, the decrease in AglB mRNA levels was 10- to 20-fold greater than the observed reduction in AglF, AglG or AglI mRNA levels.
A very different picture was obtained when real-time RT-PCR was performed with cells grown in the presence of reduced or elevated salt concentrations. In the case of cells grown in 1.75 M NaCl-containing medium, statistically significant (P < 0.01) increases in AglF, AglG and AglI mRNA levels were detected, while in cells grown in 4.8 M NaCl-containing medium, statistically significant (P < 0.01) increases in AglF and AglI mRNA levels were noted, in both cases relative to those levels measured in cells grown to mid-exponential phase in medium containing 3.5 M NaCl. The observed increases in transcription were higher in the case of cells grown in low-salt conditions. In both low- and high-salt growth conditions, the increase in AglB mRNA levels was not statistically significant.
Thus, real-time RT-PCR reveals that changes in the transcription of aglB, aglF, aglG and aglI transpire in a co-ordinated manner in the face of different growth conditions, although the nature of such changes depends on the conditions experienced, possibly reflecting an adaptive role of N-glycosylation in H. volcanii.