Abstract
All obligate intracellular pathogens or symbionts of eukaryotes lack glycogen metabolism. Most members of the Chlamydiales order are exceptions to this rule as they contain the classical GlgA-GlgC-dependent pathway of glycogen metabolism that relies on the ADP-Glucose substrate. We surveyed the diversity of Chlamydiales and found glycogen metabolism to be universally present with the important exception of Criblamydiaceae and Waddliaceae families that had been previously reported to lack an active pathway. However, we now find elements of the more recently described GlgE maltose-1-P-dependent pathway in several protist-infecting Chlamydiales. In the case of Waddliaceae and Criblamydiaceae, the substitution of the classical pathway by this recently proposed GlgE pathway was essentially complete as evidenced by the loss of both GlgA and GlgC. Biochemical analysis of recombinant proteins expressed from Waddlia chondrophila and Estrella lausannensis established that both enzymes do polymerize glycogen from trehalose through the production of maltose-1-P by TreS-Mak and its incorporation into glycogen’s outer chains by GlgE. Unlike Mycobacteriaceae where GlgE-dependent polymerization is produced from both bacterial ADP-Glc and trehalose, glycogen synthesis seems to be entirely dependent on host supplied UDP-Glc and Glucose-6-P or on host supplied trehalose and maltooligosaccharides. These results are discussed in the light of a possible effector nature of these enzymes, of the chlamydial host specificity and of a possible function of glycogen in extracellular survival and infectivity of the chlamydial elementary bodies. They underline that contrarily to all other obligate intracellular bacteria, glycogen metabolism is indeed central to chlamydial replication and maintenance.
Introduction
Chlamydiae forms with Planctomycetes and Verrucomicrobia phyla a very ancient monophyletic group of bacteria known as PVC, which has been recently enriched with additional phyla [1]. The Chlamydiales order groups the members of the Chlamydiaceae family that includes etiological agents of humans and animals infectious diseases and at least eight additional families commonly named “chlamydia-related bacteria or “environmental” chlamydia [2, 3].
The hallmark of Chlamydiales consists in an obligate intracellular lifestyle due to a significant genome reduction and biphasic development, which includes two major morphological and physiological distinct stages: the elementary body (EB), a non-dividing and infectious form adapted to extracellular survival and the reticulate body (RB), a replicating form located within a membrane surrounded inclusion (for review [4]). Following entry into a susceptible cell the EBs differentiate into RBs within the inclusion. During the intracellular stage, RBs secrete many effector proteins through the type III secretion system and express a wide range of transporters in order to manipulate host metabolism and uptake all the metabolites required for its replication. At the end of the infection cycle, RBs differentiate back into EBs before they are released into the environment [5, 6].
Glycogen metabolism loss appears to be a universal feature of the reductive genome evolution experienced by most if not all obligate intracellular bacterial pathogens or symbionts including Anaplasma spp., Ehrlichia spp., Wolbachia spp., Rickettsia spp. (alpha-proteobacteria), Buchnera sp. Coxiella sp. (gamma-proteobacteria), or Mycobacterium leprae (Terrabacteria) [7, 8]. Despite the more advanced genome reduction experienced by the animal-specific Chlamydiaceae family (0.9 Mpb) in comparison to other protist-infecting Chlamydiales (2 to 2.5 Mpb), the glycogen metabolism pathway appears surprisingly preserved [7]. This includes the three-enzymatic activities required for glycogen biosynthesis: GlgC, GlgA and GlgB. ADP-glucose pyrophosphorylase (GlgC) activity that controls the synthesis and level of nucleotide-sugar, ADP-glucose, dedicated solely to glycogen biosynthesis. Glycogen synthase (GlgA) belongs to the Glycosyl Transferase 5 family (GT5: CaZy classification) which polymerizes nucleotide-sugar into linear α-1,4 glucan.
GlgA activity has a dual function consisting of a primer-independent glucan synthesis and glucan elongation at the non-reducing end of preexisting polymers [9]. When the primer reaches a sufficient degree of polymerization (DP>15) to fit the catalytic site of the glycogen branching enzyme (GlgB), α-1,6 branches are introduced resulting in the appearance of two non-reducing polymer ends that may be further elongated by GlgA. The repetition of this process results in an exponential increase in the number of non-reducing ends leading to a particle with a 32-40 nm diameter [10].
Until recently, Waddlia chondrophila (family Waddliaceae) as well as all members of Criblamydiaceae could be considered as important exceptions to the universal requirement of Chlamydiales for glycogen synthesis. Indeed, genome analysis indicated that ad minima the glgC gene was absent from all these bacteria [11–13] and that the function of GlgA was possibly also impaired. Consequently, based on the absence of glycogen reported for all knockout glgC mutants in bacteria and plants it was believed that W. chondrophila was defective in glycogen synthesis [14, 15]. Using transmission electron microscope analysis, we are now reporting numerous glycogen particles within the cytosol of W. chondrophila and Estrella lausannensis (family Criblamydiaceae) EBs, suggesting either another gene encodes a phylogenetically distant protein that overlaps the GlgC activity or an alternative glycogen pathway takes place in these Chlamydiales.
The recent characterization of an alternative glycogen pathway, the so-called GlgE-pathway, in Mycobacterium tuberculosis and streptomycetes prompted us to probe chlamydial genomes with homolog genes involved in this pathway [16, 17]. At variance with the nucleotide-sugar based GlgC-pathway, the GlgE-pathway consists of the polymerization of α-1,4 glucan chains from maltose 1-P. In Mycobacteria, the latter is produced either from the condensation of glucose-1-P and ADP-glucose catalyzed by a glycosyl transferase called GlgM or from the interconversion of trehalose (α-α−1,1 linked D-glucose) to maltose followed by the phosphorylation of maltose, which are catalyzed by trehalose synthase (TreS) and maltose kinase (Mak) activities, respectively [16]. At the exception of Actinobacteria (i.e mycobacteria and Streptomycetes), TreS is usually fused with a maltokinase (Mak) that phosphorylates maltose into maltose-1-phosphate [18]. Subsequently, maltosyl-1-phosphate transferase (GlgE) mediates the formation of α-1,4-linked polymers by transferring the maltosyl moiety onto the non-reducing end of a growing α-1,4-glucan chain. As in the GlgC-pathway, branching enzyme (GlgB) introduces α-1,6 linkages to give rise to a highly branched α-glucan. The GlgC-pathway is found in approximately one third of the sequenced bacteria and is by far the most widespread and best studied; the GlgE pathway has been identified in 14% of the genomes of α-, β-γ-proteobacteria while 4% of bacterial genomes possess both GlgC- and GlgE-pathways [18, 19].
In order to shed light on the metabolism of storage polysaccharide in Chlamydiales, we analyzed 220 genomes (including some genomes assembled from metagenomic data) from 47 different chlamydial species that represent the bulk of currently known chlamydial diversity. A complete GlgE-pathway was identified in five chlamydial species distributed in Criblamydiaceae, Waddliaceae and Parachlamydiaceae families. In this work, we demonstrated that the GlgC-pathway is impaired in Criblamydiaceae and Waddliaceae. The complete biochemical characterization of the GlgE-pathway in Estrella lausannensis (family Criblamydiaceae) and Waddlia chondrophila (family Waddliaceae) isolated respectively from water in Spain [20, 21] and from the tissue of an abortive bovine fetus [22, 23] is reported. Thus, despite the intensive reductive genome evolution experienced by these intracellular bacteria our work shows that glycogen biosynthesis is maintained in all Chlamydiales and suggests a hitherto understudied function of storage polysaccharides and oligosaccharides in the lifecycle of all Chlamydiales.
Materials and methods
Comparative genomic analysis of glycogen metabolic pathways
In order to gain insight into Chlamydiae’s glycogen metabolism, homologs of proteins part of the glycogen pathway of E.coli and of M. tuberculosis were searched with BLASTp in 220 genomes and metagenome-derived genomes from 47 different chlamydial species available on the ChlamDB database (https://chlamdb.ch/, https://academic.oup.com/nar/article/48/D1/D526/5609527)[24]. The completeness of metagenome-derived and draft genomes was estimated with checkM based on the identification of 104 nearly universal bacterial marker genes [3]. The species phylogeny has also been retrieved from ChlamDB website.
Microscopy analysis
Fresh cultures of Acanthamoeba castellanii grown in 10 mL YPG (Yeast extract, peptone, glucose) were infected with one-week-old 5 µm-filtered suspension of E. lausannensis or W. chondrophila (105 cells.mL-1), as previously reported [25].Samples of time course infection experiments were harvested at 0, 7, 16 and 24 hours post-infection by centrifuging the infected A. castellanii cultures at 116 g. Pellets were then fixed with 1 mL of 3 % glutaraldehyde for four hours at 4°C and prepared as described previously [26].
Glycogen synthase and glycogen branching enzyme activities
Fused glgAglgB genes of E. lausannensis and W. chondrophila were amplified using primer couples harboring attB sites as described in the S1 table. PCR products were then cloned in the pET15b (Novagen) plasmid modified compatible with GatewayTM cloning strategy. The expression of his-tagged recombinant protein GlgA-GlgB was performed in the derivative BW25113 strain impaired in the endogenous glycogen synthase activity (ΔglgA). Glycogen synthases assay and zymogram analysis have been conducted as described previously [27]. The nucleotide-sugar specificity of glycogen synthase was carried out by following the incorporation of 14C-Glc of radiolabelled ADP-14C-[U]-glc or UDP-14C-[U]-glc into glycogen particles during one hour at 30°C.
Phylogeny analysis
Homologous sequences of TreS-Mak and GlgE were carried out by BLAST against the nr database from NCBI with respectively WP_098038072.1 and WP_098038073.1 sequences of Estrella lausannensis. We retrieved the top 2000 homologs with an E-value cut off ≤ 10 -5 and aligned them using MAFFT [28] with the fast alignment settings. Block selection was then performed using BMGE [29] with a block size of 4 and the BLOSUM30 similarity matrix. Preliminary trees were generated using Fasttree [30] and ‘dereplication’ was applied to robustly supported monophyletic clades using TreeTrimmer [31] in order to reduce sequence redundancy. For each protein, the final set of sequences was selected manually. Proteins were re-aligned with MUSCLE [32] block selection was carried out using BMGE with a block size of four and the matrix BLOSUM30, and trees were generated using Phylobayes [33] under the catfix C20 + Poisson model with the two chains stopped when convergence was reached (maxdiff<0.1) after at least 500 cycles, discarding 100 burn-in trees. Bootstrap support values were estimated from 100 replicates using IQ-TREE [34] under the LG4X model and mapped onto the Bayesian tree.
GlgE and TreS-Mak expressions
glgE and treS-mak genes were amplified from the genomic DNA of E. lausannensis and W. chondrophila by the primers F_glgE_EL/R_glgE_EL, F_glgE_WC/R_glgE_WC and F_treS-mak_EL/R_treS-mak_EL (S1 table). The PCR products were cloned in the expression vector pET15b (Novagene) or VCC1 (P15A replicon). The resulting plasmids pET-GlgE-WC, pET-GlgE-EL, VCC1-treS-mak-EL were transferred to E. coli Rosetta™ (DE3; pRARE) or BL21-AI™. The expression of his-tagged proteins was induced in Lysogeny Broth (LB) or Terrific Broth (TB) by the addition of 0.5 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) or 1 mM IPTG/0.2 % L-arabinose at the mid-logarithmic phase growth (A600=0.5), or by using auto inducible medium as described in Fox and Blommel [35]. After 18h at 30°C, cells were harvested at 4000g at 4°C during 15 minutes. Cell pellets were stored at −80°C until purification step on Ni 2+ affinity column.
Protein purification
Cell pellets from 100 mL of culture medium were resuspended in 1.5 mL of cold buffer (25 mM Tris-acetate, pH 7.5). After sonication (three times 30 s), proteins were purified on Ni 2+ affinity column (Roth) equilibrated with washing buffer (300 mM NaCl, 50 mM sodium acetate and 60 mM imidazole, pH 7) and eluted with a similar buffer containing 250 mM imidazole. Purification steps were followed by SDS-PAGE and purified enzymes quantified by Bradford method (Bio-Rad).
Evidence of GlgE-like activity by thin layer chromatography
Maltosyl transferase activities of GlgE proteins of E. lausannensis and W. chondrophila were first evidenced by incubating the purified recombinant enzymes overnight at 30°C with 10 mg.mL-1 glycogen from rabbit liver (Sigma-Aldrich) and 20 mM inorganic phosphate in 20 mM Tris-HCl buffer (pH 6.8). The reaction products were separated on thin layer chromatography Silica gel 60 W (Merck) using the solvent system butanol/ethanol/water (5/4/3 v/v/v) before spraying orcinol (0.2%)-sulfuric (20%) solution to visualize carbohydrates.
Maltose-1-phosphate purification
Maltose-1-phosphate was purified from 20 mL of enzymatic reaction with GlgE-EL (1mg GlgE-EL, 10 mg.mL-1 potato amylopectin and 20 mM orthophosphate in 20 mM TRIS/HCl pH 6.8, at 30°C, overnight) with several steps. First, size exclusion chromatography on TSK-HW 50 (Toyopearl, 48 x 2.3 cm, flow rate of 1 mL.min-1) equilibrated with 1% ammonium acetate was used to remove glycogen. Maltose-1-phosphate was separated from orthophosphate by anion exchange chromatography using Dowex 1 × 8 100-200 mesh (Bio-Rad, 28 x 1.6 cm, acetate form, flow rate of 0.75 mL.min-1) in 0.5 M potassium acetate pH 5, then neutralized with ammoniac, and purified from remaining salts using Dowex 50 W X 8 50-100 mesh (Bio-Rad, 10 x 1 cm, H+ form) equilibrated with water. Around 10 mg of maltose-1-phosphate were recovered from a reaction mixture of 20 mL, with a yield of approximately 5%. The end product was used for mass spectrometry and NMR analysis. Maltose-1-phosphate was also produced by incubating overnight at 30°C recombinant TreS-Mak protein from E. lausannensis with 20 mM ATP, 20 mM maltose, 10 mM MnCl2, 125 mM imidazole and 150 mM NaCl. After an anion exchange chromatography as described above, maltose-1-phosphate was purified from remaining maltose and salts using Membra-cell MC30 dialysis membrane against ultrapure water. This purification procedure leads to a better yield (around 8%).
Proton-NMR analysis of maltose-1-phosphate
Sample was solubilized in D2O and placed into a 5mm tube. Spectra were recorded on 9.4T spectrometer (1H resonated at 400.33 MHz and 31P at 162.10MHz) at 300K with a 5 mm TXI probehead. Used sequences were extracted from Bruker library sequence. Delays and pulses were optimized for this sample.
MALDI-TOF MS Analysis
P-maltose was analyzed by a MALDI-QIT-TOF Shimadzu AXIMA Resonance mass spectrometer (Shimadzu Europe; Manchester, UK) in the positive mode. The sample was suspended in 20 µL of water. 0.5 µL sample was mixed with 0.5 µL of DHB matrix on a 384-well MALDI plate. DHB matrix solution was prepared by dissolving 10 mg of DHB in 1 mL of a 1:1 solution of water and acetonitrile. The low mode 300 (mass range m/z 250-1300) was used and laser power was set to 100 for 2 shots each in 200 locations per spot.
Kinetic parameters of GlgE and TreS-Mak of E. lausannensis
GlgE activity was monitored quantitatively in the elongation direction by the release of orthophosphate using the Malachite Green Assay Kit (Sigma-Aldrich) following the manufacturers’ instructions. The concentration of released free phosphate was estimated from a standard curve, monitoring the absorbance at 620 nm with Epoch microplate spectrophotometer (Biotek). Kinetic parameters of GlgE-EL were determined in triplicates at 30°C in 15 mM Tris/HCl buffer at pH 6.8.
Saturation plots for maltose-1-phosphate were obtained with 10 mM of maltoheptaose or 10 mg.mL-1 of glycogen from bovine liver (Sigma-Aldrich) whereas 2 mM maltose-1-phosphate were used to get saturation plots for maltoheptaose and glycogen from bovine liver. Optimal temperature and pH were assayed with 1 mM maltose-1-phosphate and 5 mM maltoheptaose, respectively in 25 mM Tris/HCl pH 6,8 and at 30°C. Temperature was tested in the range of 15°C to 45°C and pH between 3 and 8.8 with different buffers: 25 mM sodium acetate at pH 3.7, 4.8 and 5.2; 25 mM sodium citrate at pH 3, 4, 5 and 6; 25 mM Tris/HCl at pH 6.8, 7, 7.5, 7.8, 8 and 8,8.
The TreS activity domain of TreS-Mak protein was monitored following the conversion of trehalose into maltose and glucose using Epoch spectrophotometer (Biotek). 15µL of reaction sample were incubated 30 min at 58°C with 30 µL of 100 mM sodium citrate pH 4.6 containing whether 0.4 U of amyloglucosidase from Aspergillus niger (Megazyme) or no amyloglucosidase to then quantify the amount of free glucose and maltose after addition of 100 µL of a buffer (500 mM triethanolamine hydrochloride, 3.4 mM NADP+, 5 mM MgSO4 and 10 mM ATP, pH 7.8). The increase in absorbance at 340 nm after the supplementary addition of 1.2 U of hexokinase and 0.6 U of glucose-6-phosphate dehydrogenase (Megazyme) allowed us to estimate the amount of glucose units from a standard curve. Unless otherwise stated, enzymatic reactions were performed at 30°C, pH 8, with 125 mM imidazole, 150 mM NaCl, 200 mM trehalose and 1 mM MnCl2.
The maltokinase activity of TreS-Mak protein was monitored following the amount of nucleoside bi-phosphate released. 20 µL of reaction mixtures were added to 80 µl of pyruvate kinase buffer (75 mM Tris/HCl pH 8.8, 75 mM KCl, 75 mM MgSO4, 2 mM phosphoenolpyruvate, 0.45 mM reduced NADH). The amount of nucleoside diphosphate was estimated from a standard curve of ADP, measuring the decrease in absorbance at 340 nm 30 min after addition of 5 U of L-lactic dehydrogenase (from rabbit muscle, Sigma-Aldrich) and 4 U of pyruvate kinase (from rabbit muscle, Sigma-Aldrich). If not stated otherwise, enzymatic reactions were performed at 30°C, pH 8, with 125 mM imidazole, 150 mM NaCl, 20 mM maltose, 20 mM ATP (or other nucleoside triphosphate) and 10 mM MnCl2.
Chain length distribution analyses
GlgE activity was qualitatively monitored using fluorophore-assisted carbohydrates electrophoresis (FACE). Reactions with 5 mM of malto-oligosaccharides from glucose to maltoheptaose, 1.6 mM M1P and a specific activity of 3.5 nmol orthophosphate produced per minute for GlgE_EL and 1.4 nmol/min for GlgE_WC, were performed in a 100 µL volume at 30°C during 1h and 16h. Reactions were stopped at 95°C for 5 min and supernatants recovered after centrifugation. Samples were dried and resolubilized in 2 µL 1 M sodium cyanoborohydride (Sigma-Aldrich) in THF (tetrahydrofurane) and 2 µL 200 mM ATPS (8-aminopyrene-1,3,6-trisulfonic acid trisodium salt, Sigma-Aldrich) in 15% acetic acid (v/v). Samples were then incubated overnight at 42°C. After addition of 46 µL ultrapure water, samples were again diluted 300 times in ultrapure water prior to injection in a Beckman Coulter PA800-plus Pharmaceutical Analysis System equipped with a laser-induced fluorescence detector. Electrophoresis was performed in a silicon capillary column (inner diameter: 50 µm; outer diameter: 360 µm; length: 60 cm) rinsed and coated with carbohydrate separation gel buffer-N (Beckman Coulter) diluted 3 times in ultrapure water before injection (7 s at 10 kV). Migration was performed at 10 kV during 1 h.
1 mg glycogen from bovine liver (Sigma) and de novo polysaccharide produced from overnight incubation of 2 mg maltose-1-phosphate with 30 µg GlgE_EL and 200 µg GlgB_WC were purified by size exclusion chromatography on TSK-HW 50 (Toyopearl, 48 x 2.3 cm, flow rate of 0.5 mL/min) equilibrated with 1% ammonium acetate. Remaining maltose-1-phosphate was dephosphorylated with 10 U of alkaline phosphatase (Sigma-Aldrich) overnight at 30°C and samples were dialyzed using Membra-Cel MC30 dialysis membrane against ultrapure water. The chain length distribution of samples was then analysed following protocol described just above, with slight differences. Prior to APTS labelling, samples were debranched overnight at 42°C in 50 mM sodium acetate pH 4.8 by 2 U of isoamylase from Pseudomonas sp. (Megazyme) and 3.5 U of pullulanase M1 from Klebsiella planticola (Megazyme), then desalted with AG® 501X8(D) Mixed Bed Resin. Labelled samples were diluted 10 times in ultrapure water before injection.
Zymogram analysis
7.5% acrylamide-bisacrylamide native gels containing 0.3% glycogen from bovine liver (Sigma-Aldrich) or 0.3% potato starch (w/v) were loaded with 2 µg of crude protein extract or purified recombinant enzyme. Migration was performed in ice-cold TRIS (25 mM) glycine (192 mM) DTT (1 mM) buffer, during 2 h at 120 V and 15 mA per gel, using MiniProtean II (Biorad) electrophoresis system. Gels were then incubated overnight, at room temperature and under agitation, in 10 mL Tris (25 mM) acetate pH 7.5 DTT (0.5 mM), supplemented when stated, with 1 mM maltose-1-phosphate or 20 mM orthophosphate. Gels were rinsed 3 times with ultrapure water prior staining with iodine solution (1% KI, 0.1% I2).
Determination of the apparent molecular weight of GlgE and TreS-Mak
The apparent molecular weight of recombinant GlgE_EL and TreS-Mak_EL were determined using native PAGE and gel filtration. For native PAGE, 5%, 7.5%, 10% and 15% acrylamide: bisacrylamide (37.5 : 1) gels (20 cm x 18.5 cm x 1 mm) were loaded with 6 µg of protein of interest and some standard proteins of known mass: 15 µg carbonic anhydrase (29 kDa), 20 µg ovalbumin (43/86 kDa), 15 µg BSA (66.5/133/266/532 kDa), 15 µg conalbumin (75 kDa), 1.5 µg ferritin (440 kDa) and 25 µg thyroglobuline (669 kDa). Log10 of migration coefficient was plotted against the acrylamide concentration in the gel. Negative slopes were then plotted against molecular weights of standard proteins and the apparent molecular weight of proteins of interest was determined using slope equation. Gel permeation chromatography, Sepharose™ 6 10/300 GL resin (30 cm x 1 cm; GE Healthcare) was equilibrated in PBS buffer (10 mM orthophosphate, 140 mM NaCl, pH 7.4) at 4°C and with a flow rate of 0.3 mL/min. Void volume was determined using Blue Dextran 2000.
Standard proteins used were ribonuclease A (13.7 kDa, 3 mg.mL-1), ovalbumin (43 kDa, 4 mg.mL-1), aldolase (158 kDa, 4 mg.mL-1), ferritin (440 kDa, 0.3 mg.mL-1) and thyroglobuline (669 kDa, 5 mg.mL-1). All standard proteins used were from GE’s Gel Filtration Low Molecular Weight Kit and GE’S Gel Filtration High Molecular Weight Kit (GE Healthcare), except for the BSA (Sigma-Aldrich).
Results
Two different glycogen metabolic pathways identified in the Chlamydiae phylum
To gain insight into Chlamydiae’s glycogen metabolism, we analyzed 220 genomes from 47 different chlamydial species. As illustrated in figure 1A, the synthesis of linear chains of α-1,4 glucose involves both ADP-glucose pyrophosphorylase (GlgC) and glycogen synthase (GlgA) activities in the GlgC-pathway while GlgE-pathway relies on trehalose synthase (TreS), maltokinase (Mak) and maltosyl-1 phosphate transferase (GlgE). The formation of α-1,6 linkages (i.e. branching points) and glycogen degradation are catalyzed by a set of similar enzymes in both pathways that include glycogen branching enzyme isoforms (GlgB/GlgB2) and glycogen phosphorylases isoforms (GlgP /GlgP2), glycogen debranching enzymes (GlgX) and α-1,4 glucanotransferase (MalQ). The genomic database used in this study (https://chlamdb.ch) includes genomes from both cultured and uncultured Chlamydiae species that cover the diversity of the chlamydiae phylum (figure 1B). Comparative genomics clearly underlined the high prevalence of a complete GlgC-path in most Chlamydiales, including all members of the Chlamydiaceae family, which undergoes massive genome reduction (identified by the letter “d” on figure 1B) as well as in in the most deeply branching families such as candidatus Pelagichlamycidiaceae (“a”) and candidatus Parilichlamydiaceae (“b”). We noticed that the glg genes are at least 10 kbp apart with a notable exception for glgP and glgC, which are mostly separated by one or two genes. It should be stressed out that the gaps in glycogen metabolism pathways of several uncultivated chlamydiae likely reflect the fact that many of those genomes are incomplete genomes derived from metagenomic studies (see percentages in brackets in figure 1B). Considering that the GlgC-pathway is highly conserved in nearly all sequenced genomes of the phylum, missing genes probably reflect missing data rather than gene losses. It is interesting to note that there is uncertainty about the presence of glgC gene in Candidatus Enkichlamydia genome (“j”), a complete set of glycogen metabolizing enzymes were recovered expect for gene encoding for ADP-glucose pyrophospharylase (glgC). This gene is missing from 6 independent draft genomes estimated to be 71% to 97% complete, suggesting either the loss of glgC gene or that glgC gene is located in a particular genomic region (e.g. next to repeated sequences) that systematically led to its absence from genome assemblies. Another unexpected result concerns both Waddliaceae (“l”) and Criblamydiaceae (“m”) families that encompass Waddlia chondrophila, Estrella lausannensis and Criblamydia sequanensis species. Genomic rearrangements caused a sequence of events leading to (i) the deletion of both glgC and glgP genes (ii) the fusion of glgA with glgB gene (iii), the insertion of glgP2 gene encoding glycogen phosphorylase isoform at the vicinity of malQ gene. It should be stressed out that a homolog of glgP2 gene has also been identified on the plasmids of S. nevegensis and P. naegleriophila. In W. chondrophila, another insertion of glgP2 occurred downstream to the GlgE operon, which may be correlated with partial deletion of glgP2 at the vicinity of malQ (Figure 1B). The parsimonious interpretation of glgC and glgP deletion and glgAglgB fusion is that a single deletion event led to the loss of DNA fragment bearing glgP and glgC genes between glgA and glgB. Despite many variations, we did not observe such configuration in the chlamydial genome analyzed (S2 Table). More remarkably, genomic rearrangements are associated with a novel glycogen pathway based on GlgE operon described in mycobacteria and also observed in Prototochlamydia naegleriophila and Protochlamydia phocaeensis (syn. Parachlamydia C2). All three genes are clustered in the classical unfused glgE-treSmak-glgB2 operon arrangement in Waddliaceae and Criblamydiaceae, while the glgB2 gene is missing in the Parachlamydiaceae operons (Figure 1B). The occurrence of GlgE pathway restricted to Parachlamydiaceae, Waddliaceae and Criblamydiaceae families arises questions about its origin in Chlamydiales. To get some insight on this issue, phylogenetic trees of TreS-Mak and GlgE have been inferred using the phylobayes method (Figure 2). The GlgE phylogeny shows that even if the Chlamydiae sequences are split into two with W. chondrophila on one side and the other sequences on the other side, which reflects likely lateral gene transfer events with other bacteria, chlamydial glgE sequences might still be monophyletic since the only strongly supported node (marked as red star) with a posterior probability (pp) higher than 0.95 (pp = 0.99) unifies all chlamydiae sequences, which has also been confirmed using LG4X model (Data not shown) (Figure 2A). The phylogeny analysis highlights that GlgE sequences can be classified into classes I and II, comprising Chlamydiales and Actinomycetales (i.e. mycobacteria, Streptomycetes), respectively. For Tres-Mak phylogeny, chlamydial Tres-Mak sequences cluster together, suggesting a common origin, however with a low statistical support (pp=0.93). Although the origin of GlgE operon cannot be pinpointed in our phylogenetic analysis, conceivable scenarios are that either i) GlgE operon reflects vestigial metabolic function of the ancestral chlamydiae and then has been lost in most families or ii) this operon was acquired by lateral gene transfer event from a member of PVC phylum by the common ancestor of Parachlamydiaceae, Waddliaceae and Criblamydiaceae families.
Classical GlgC-pathway is not functional in E. lausannensis and W. chondrophila
To further investigate whether his-tagged recombinant proteins GlgA-GlgB of E. lausannensis and W. chondrophila are functional, glycogen synthase activities at the N-terminus domain were assayed by measuring the incorporation of labeled 14C-glucosyl moiety from ADP- or UDP-14C-glucose onto glycogen and by performing a specific non-denaturing PAGE or zymogram to visualize glycogen synthase activities. After separation on native-PAGE containing glycogen, recombinant proteins were incubated in the presence of 1.2 mM ADP-glucose or UDP-glucose, glycogen synthase activities are visualized as dark activity bands after soaking gels in iodine solution (Figure 3). Enzymatic assays and zymogram analyses show that the glycogen synthase domain of the chimeric GlgA-GlgB of W. chondrophila (hereafter GlgA-GlgB-WC) is functional but highly specific for ADP-glucose (0.70 nmol of incorporated glucose. min-1.mg-1) and has little to no activity using UDP-glucose as substrate. As predicted, the activity of the truncated glycogen synthase in E. lausannensis was not detected on activity gels or during enzymatic assays (S1A Figure).
We further investigated whether the branching activity domain at the carboxyl terminal of chimeric protein GlgA-GlgB of W. chondrophila (GlgA-GlgB-WC) was functional. To check this, the same chimeric GlgA-GlgB-WC sample previously analyzed was incubated with ADP-glucose (3 mM) and maltoheptaose (10 mg.mL-1) overnight. Subsequently, the appearance of branching point (i.e α-1,6 linkages) onto growing linear glucans can be specifically observed by the resonance of protons onto carbon 6 at 4.9 ppm using proton-NMR analysis. However as depicted on S1C figure, we did not observe any signal, suggesting that branching enzyme activity domain is not functional despite an active glycogen synthase domain. This result is consistent with several reports indicating that the amino-acid length at the N-terminus of branching enzyme affects its catalytic properties [36, 37]. In regards to this information, the glycogen synthase domain extension located at the N-terminus prevents probably the branching enzyme activity of GlgA-GlgB. Thus α-1,6 linkages or branching points are likely to be the result of GlgB2 isoform activity found in both instances. Altogether, these data strongly suggest that the classical GlgC-pathway is not functional in both Waddliaceae and Criblamydiaceae families.
GlgE-like genes of E. lausannensis and W. chondrophila encode α-maltose 1-phosphate: 1,4-α-D-glucan 4-α-D-maltosyl transferase
Based on phylogenetic analysis of GlgE, both GlgE of mycobacteria (Actinobacteria) and Chlamydiales are phylogenetically distant (Figure 2A). GlgE of M. tuberculosis displays 43% to 40% of identity with GlgE-like sequences of E. lausannensis and W. chondrophila, respectively. Because GlgE activity belongs to the large and diversified Glycosyl Hydrolase 13 family consisting of carbohydrate active enzymes with quite diverse activities such as α-amylases, branching enzymes, debranching enzymes [38], we undertook to demonstrate that these enzymes displayed catalytic properties similar to those previously described for GlgE of mycobacteria. Histidine-tagged recombinant proteins of GlgE of Estrella lausannensis (hereafter GlgE-EL) and Waddlia chondrophila (hereafter GlgE-WC) were expressed and further characterized (S2 Figure). As described in previous studies, GlgE of Mycobacteria mediates the reversible reaction consisting into the release of maltose-1-phosphate in the presence of orthophosphate and α-glucan polysaccharide. Both GlgE-EL and GlgE-WC were incubated in presence of glycogen from rabbit liver and orthophosphate. After overnight incubation, reaction products were analyzed on thin layer chromatography and sprayed with oricinol-sulfuric acid (Figure 4A). A fast migration product capable of interacting with oricinol sulfuric acid was clearly synthesized in crude extract (CE), in washing # 3 (W3) and in the purified enzyme fraction (E1) of GlgE-EL sample. A barely visible product is only observed in the purified fraction (E1) of GlgE-WC. To further characterize this material, time course analysis of phosphatase alkaline (PAL) treatment was performed on the reaction product suspected to be M1P obtained from sample E1 of GlgE-EL. After 180 min of incubation, the initial product is completely converted into a compound with a similar mobility than maltose (DP2) (Figure 4B). The compound produced by GlgE-EL in presence of glycogen and orthophosphate were further purified through different chromatography steps and subjected to mass spectrometry and proton-NMR analyses (Figures 4C and 4D). The combination of these approaches confirms that GlgE of E. lausannensis as well as W. chondrophila (S3 Figure) catalyzes the formation of a compound of 422 Da corresponding to α-maltose-1-phosphate. In order to carry out enzymatic characterization of GlgE activities, identical purification processes were scaled up to purify enough M1P, free of inorganic phosphate and glucan.
Kinetic parameters of GlgE activity of E. lausannensis in the biosynthetic direction
Because the his-tagged recombinant GlgE-WC expresses very poorly and the specific activity of GlgE-WC was ten times lower than GlgE-EL, kinetic parameters were determined in the synthesis direction i.e. the transfer (amount) of maltosyl moieties onto non-reducing ends of glucan chains, exclusively for GlgE-EL. Transfer reactions are associated with the release of inorganic phosphate that can be easily monitored through sensitive malachite green assay. Thus, under variable M1P concentrations and using fixed concentrations of glycogen or maltoheptaose, the GlgE-EL activity displays allosteric behavior indicating a positive cooperativity, which has been corroborated with Hill coefficients that were above 1 (Figures 5 A and 5 B). In agreement with this, the molecular weight of native GlgE-EL determined either by size exclusion chromatography or by native-PAGE containing different acrylamide concentrations (5%; 7.5 %; 10 % and 12.5 %) indicates an apparent molecular weight of 140 to 180 kD respectively corresponding to the formation of dimer species while no monomer species of 75 kD were observed (Figure 5E). The enzyme exhibited S0.5 values for M1P that vary from 0.16±0.01 mM to 0.33±0.02 mM if DP7 and glycogen are glucan acceptors, respectively. However, using M1P at saturating concentration, GlgE-EL displays Michaelis kinetics (nH close to 1) indicating a non-cooperative reaction (figures 5C and 5 D). In such experimental conditions, the apparent Km values for glycogen and DP7, 2.5 ± 0.2 mg.mL-1 and 3.1 ± 0.2 mM, respectively were similar to the apparent Km value of glycogen synthase (GlgA) that synthesizes α-1,4 linkages from ADP-glucose [39].
De novo glycogen synthesis: GlgE activity enables the initiation and elongation of glucan
Contrarily to eukaryotic glycogen synthase, prokaryotic glycogen synthase (GlgA) does not require the presence of a short α-1,4 glucan or primer to initiate glycogen biosynthesis [9]. In absence of GlgA and GlgC activity in E. lausannensis and in the absence of GlgC and thus of ADP-glucose supply in W. chondrophila, this raised the question of the ability of GlgE activities to substitute for GlgA with respect to the priming of glycogen biosynthesis. To establish whether GlgE activities are able to prime glucan synthesis, both his-tagged GlgE-EL (3.51 nmol of Pi released.min-1) and GlgE-WC (1.38 nmol of Pi released.min-1) were incubated with 1.6 mM M1P and in the presence of 5mM of various glucan chains with a degree of polymerization (DP) of 1 to 7. Identical incubation experiments were conducted with GlgE recombinant proteins except M1P was omitted in order to appreciate α-1,4 glucanotransferase or disproportionnating activity (Figure 6 and S4, S5 Figures). After incubation, the reduced-ends of glucan chains were labeled with fluorescent charged probe (APTS) and separated according to their degree of polymerization by capillary electrophoresis. We noticed that the C1 phosphate group prevented the labeling of M1P with fluorescent probe.
Nevertheless, the level of maltose released from M1P due to the spontaneous dephosphorylation during the experiment was appreciated by performing incubations with denatured enzymes (Figures 6A and 6H). Incubation experiments show that both GlgE activities possess either an α-1,4 glucanotransferase or maltosyltransferase activities depending on the presence of M1P. When M1P is omitted, GlgE activities harbor an α-1,4 glucanotransferase activity exclusively with glucans composed of six or seven glucose units (DP6 or DP7). Interestingly, after one hour or overnight incubation, DP6 or DP7 are disproportionated with one or two maltosyl moieties leading to the release of shorter (DPn-2) and longer glucans (DPn+2) (Figures 6G and 6N and S4, S5 Figures). The limited number of transfer reactions emphasizes probably a side reaction of GlgE activities. The α-glucanotransferase activity can be also appreciated on native-PAGE containing glycogen. Chain length modification of external glucan chains of glycogen results in increase of iodine interactions visualized as brownish activity band (S6A Figure). After one hour of incubation (S4, S5 Figures), both GlgE activities enable the transfer the maltosyl moiety of M1P onto the glucan primer with a DP ≥ 3 (S4, S5 Figures). Interestingly, for a longer period of incubation time, both GlgE activities switch to either a processive or a distributive activity mode depending on the initial degree of polymerization of glucan primer. For instance, in the presence of maltose (DP2) or maltotriose (DP3) both GlgE-EL and GlgE-WC undergo processive elongation activities, which consist of the synthesis of very long glucan chains, up to 32 glucose residues. In contrast, when both GlgE activities are incubated in presence of glucan primers with DP ≥4, the latter add and immediately release a glucan primer (DP) with an increment of two glucose moieties (DPn+2) that leads to distributive elongation behavior. The mechanism underlying the switch between processive and distributive elongation activities reflects probably a competition of glucan primers for the glucan binding site in the vicinity of the catalytic domain. Thus, we can hypothesize that the low affinity of short glucan primers (DP<4) for glucan binding site favors probably iterative transferase reactions onto the same acceptor glucan (i.e. processive mode) resulting in the synthesis of long glucan chains whereas glucan primers with DP≥ 4 compete strongly for the binding site leading to a distributive mode. The discrepancy between GlgE-EL and GlgE-WC to synthesize long glucan chains in the absence (Figures 6B, 6I) or in the presence of glucose (DP1) (Figures 6C, 6J) might be explained by a higher amount of free maltose observed in denatured GlgE-WC samples (Figure 6A) by comparison to denatured GlgE-EL samples (Figure 6H). Despite having taken all precautious (same M1P preparation, buffer pH7), spontaneous dephosphorylation of M1P occurred more significantly in GlgE-WC samples. We therefore conclude that initial traces of maltose in GlgE-WC samples facilitate the synthesis of long glucan chains in the absence (Figure 6B) or in the presence of glucose (DP1) (Figure 6C). To test this hypothesis, crude extract (CE) and purified GlgE proteins (E1) of E. lausannensis were loaded onto non-denaturing polyacrylamide electrophoresis (native-PAGE). After migration, slices of polyacrylamide gel were incubated overnight in buffers containing 0 mM (control) or 2 mM M1P (Figure 7A). The synthesis of long glucan chains with DP> 15 (minimum number of glucose units required for detection through interaction with iodine molecules) are detected by soaking the gel in iodine solution. As depicted in Figure 7A, the synthesis of glucan chains catalyzed by GlgE-EL appears exclusively as dark-blue activity bands inside native-PAGE incubated with 2 mM M1P and not in the absence of M1P. Altogether, these results suggest that GlgE activities are able to synthesize de novo a sufficient amount of long linear glucans from maltose-1-phosphate. We cannot exclude the role of maltose in the initiation process of glucan synthesis as glucan acceptor since spontaneous dephosphorylation of M1P is unavoidable. We further carried out a series of experiments that consisted to synthesize in vitro high molecular branched glucans by incubating both recombinant glycogen branching enzyme of W. chondrophila (GlgB-WC: S6B Figure) and GlgE-EL in the presence of M1P. After overnight incubation, the appearance of α-1,6 linkages or branching points were directly measured by subjecting incubation product on proton-NMR analysis (Figure 7B). In comparison with M1P and glycogen as controls, proton-NMR spectrum of incubation products show a typical profile of glycogen-like with signals at 5.6 ppm and 4.9 ppm of proton involved in a-1,4 and a-1,6 linkages.
This branched polysaccharide material was further purified and incubated with a commercial isoamylase type debranching enzyme (Megazyme) that cleaves off α-1,6 linkages or branching points. Released linear glucan chains were labeled with APTS and separated according to the degree of polymerization by capillary electrophoresis. The chain length distribution (CLD) of synthesized polysaccharides (Figure 7C) was compared with glycogen from rabbit liver (Figure 7E). As control, the amounts of free linear glucans were estimated by analyzing the APTS-labeled samples not incubated with commercial debranching enzyme (Figures 7D and 7F). In absence of significant amount of free glucan chains (Figure 7D), the in vitro synthesized polysaccharide harbors a typical CLD similar to animal glycogen with monomodal distribution and maltohexaose (DP6) as most abundant glucan chains. Altogether, these results confirm that GlgE activities display an in vitro function similar to that of glycogen synthase (GlgA) for initiating and elongating the growing glycogen particles.
Expression of bifunctional TreS-Mak of Estrella lausannensis
To our knowledge, the characterization of the bifunctional TreS-Mak activity has not yet been reported in the literature. At variance to previous GlgE expression experiments, first transformation experiment in RosettaTM (DE3) E.coli strain did not yield colonies in spite of the absence of inducer (IPTG) (S7A Figure). We presumed that a basal transcription of treS-mak gene associated with a substantial intracellular amount of trehalose (estimated at 8.5 mM in E.coli cell spread on Luria-Broth agar medium [40]) lead to the synthesis of highly toxic maltose-1-phosphate. This encouraging result prompted us to perform expression of TreS-Mak protein in BL21-AI strain. The his-tagged TreS-Mak protein purified on nickel columns displays a molecular weight of 115 kDa on SDS-PAGE (S7B Figure) while in solution recombinant TreS-Mak formed a homodimer with an apparent molecular weight of 256 kDa as analyzed by sepharose 6 column chromatography (S7C Figure). This contrast with the hetero-octameric complex of 4TreS-4 Mak (≈490 kDa) observed in Mycobacterium smegmatis in which homotetramers of TreS forms a platform to recruit dimers of Mak via specific interaction domain [41, 42].
We first confirmed that the N-terminus TreS domain is functional by measuring the interconversion of trehalose into maltose. The amount of maltose was enzymatically quantified using the amyloglucosidase assay. Previous reports indicated that TreS activities are partially or completely inhibited with 10 mM of divalent cation while a concentration of 1 mM has positive effects. The effect of Mn2+ cation on the activity of TreS domain was inferred at 200 mM of trehalose. As depicted on Figure 8A, the activity of the TreS domain increases only slightly by 1.1-fold from 0 mM to 1 mM of Mn 2+ (0.37 µmol maltose. min-1.mg-1) whereas a noticeable decrease of TreS activity (0.24 µmol maltose. min-1.mg-1) is obtained at 10 mM of Mn 2+. As reported in the literature, the TreS activity is also associated with the release of glucose during the interconversion of trehalose into maltose. Because TreS activity is fused with the Mak domain in E. lausannensis, we tested the effect of a wide range of concentration of ATP concentration on the interconversion of trehalose (Figure 8B). Although no significant effect of ATP was observed on TreS activity at 1mM (0,43 µmol maltose. min-1.mg-1), TreS activity decreased by 0.6-fold at 3 mM-10 mM ATP (0.29µmol maltose. min-1.mg-1) and dropped by 2.8-fold when the ATP concentration reaches up to 20 mM (0,15 µmol maltose. min-1.mg-1). Finally, the apparent Km value for trehalose was determined at 42.3±2.7 mM in the presence of 1 mM MnCl2 and 0 mM ATP (Figure 8C). This is consistent with the apparent Km values for trehalose (50 to 100 mM) reported in the literature for TreS activity in various species [43].
We further focused on the activity of the maltokinase domain that catalyzes the phosphorylation of maltose in presence of ATP and releases M1P and ADP. The latter was monitored enzymatically via the pyruvate kinase assay in order to express the Mak activity domain as µmol of ADP released.min-1. mg-1 of protein. Our preliminary investigations indicated that imidazole stabilizes or is required for the maltokinase activity domain (S7D Figure). Hence all incubation experiments have been conducted in the presence of 125 mM of imidazole. The pH and temperature optima were respectively determined at 30°C and pH 7 (S7E and S7F Figure). Interestingly, the activity of the Mak domain is functional within a wide range of temperature that reflects probably the temperature of free-living amoebae or animal hosts. Kinase activities are reported for their requirement in divalent cation in order to stabilize the negatively charged phosphate groups of phosphate donors such as ATP. Therefore, TreS-Mak activity was inferred in the presence of various divalent cations (Figure 8E). As expected, the recombinant TreS-Mak was strictly dependent on divalent cations, in particular, with a noticeable stimulatory effect of Mn 2+ (Figure 8D and 8E). Others tested divalent cations, like Co2+, Mg2+, Fe2+, Ca2+, Cu2+ activated the Mak activity as well, but to a lower extent, while no effect was observed in presence of Ni2+. Interestingly, at variance to Mak activity of Mycobacterium bovis, which prefers Mg 2+, the catalytic site of the Mak activity domain of TreS-Mak binds preferentially Mn 2+ over Mg 2+ [44], which is consistent with a distinct evolutionary history as depicted on figure 2B. Then, nucleotides, ATP, CTP, GTP and UTP were tested as phosphate donors by measuring the amount of M1P released (Figure 8F). The data expressed in percentage of activity show that ATP (100%), GTP (85%), UTP (70%) and to a lower extent CTP (31%) are efficient phosphate donors. Altogether, we demonstrated that TreS and Mak domains are functional in the fused protein TreS-Mak of E. lausannensis. The reversible interconversion of trehalose combined with an intracellular trehalose concentration probably below 42 mM (the intracellular trehalose concentration was estimated at 40±10 mM inside one cell of E. coli strain overexpressing OtsA/OtsB [40]) suggest that irreversible phosphorylation of maltose drives the synthesis of M1P.
Estrella lausannensis and Waddlia chondrophila accumulate glycogen particles within the cytosol of EB via GlgE-pathway
Since incubation experiments have shown that branched polysaccharide can be synthesized in the presence of maltose-1-phosphate and both GlgE and GlgB activities, this prompted us to examine the presence of glycogen particles in thin section of E. lausannensis and W.chondrophila by transmission electron microscopy. After twenty four hours post infection, thin sections of Acanthamoeba castellanii infected with both Chlamydiales (Figures 9A and 9C) and purified elementary bodies (Figures 9B and 9D) were subjected to specific glycogen staining based on the periodic acid method, which is considered to be one of the most reliable and specific methods for staining glycogen [45]. Glycogen particles appear as electron-dense particles (white head arrows) in the cytosol of elementary bodies of E. lausannensis and W. chondrophila. Interestingly, because Waddlia chondrophila infects animal cells, which do not synthesize trehalose, this suggests that trehalose must be synthesized by the bacteria itself [46]. Based on the five different trehalose pathways described in prokaryotes (for review [47]), we found that trehalose biosynthesis is limited to so-called “environmental Chlamydiae” and is not present in the Chlamydiaceae family. Among chlamydial strains with GlgE pathway, P. phocaeensis and P. neagleriophila synthesize trehalose through TreY-TreZ pathway while OtsA-OtsB pathway was found in both E. lausannensis and W. chondrophila. Importantly, otsA and otsB genes encode for trehalose-6-phosphate synthase and trehalose-6-phosphate phosphatase, respectively. OtsA activity condenses glucose-1-phosphate and UDP-glucose into trehalose-6-phosphate. However, BLAST search did not evidence the classical galU gene encoding UDP-glucose pyrophosphorylase, which synthesizes UDP-glucose from glucose-1-phosphate and UTP in both E. lausannensis and W. chondrophila, but rather a non-GalU type UDP-glucose pyrophosphorylase homolog to UGP3 of plants. This chloroplastic UDP-glucose pyrophosphorylase activity, dedicated to sulfolipid biosynthesis belongs to a set of 50 to 90 chlamydial genes identified, as lateral gene transfer, in the genomes of Archaeplastida [48]. Based on this work and taking into account the current genome analysis, we propose that glycogen metabolism pathway in W. chondrophila and E. lausannensis occur as depicted on Figure 10.
Discussion
The present study examined the glycogen metabolism pathway in Chlamydiae phylum. Unlike other obligatory intracellular bacteria, Chlamydiae have been documented to retain their capacity to synthesize and degrade the storage polysaccharide with the notable exception of the Criblamydiaceae and Waddliaceae families, for which the key enzyme of glycogen biosynthesis pathway, ADP-glucose pyrophosphorylase activity was reported missing [11, 49]. All mutants deficient in GlgC activity are associated with glycogen-less phenotypes and so far no homologous gene encoding for a GlgC-like activity has been established among prokaryotes [50, 51]. To our knowledge, only two cases have been documented for which GlgC activity has been bypassed in the classical GlgC-pathway. The ruminal bacterium Prevotella bryantii that does not encode an ADP-glucose pyrophosphorylase (glgC) gene has replaced the endogenous glgA gene with an eukaryotic UDP-glucose-dependent glycogen synthase [52, 53]. The second case reported concerns the GlgA activity of Chlamydia trachomatis which has evolved to polymerize either UDP-glucose from the host or ADP-glucose produced by GlgC activity into glucose chains [54]. In order to get some insight in Criblamydiaceae and Waddliaceae families, a survey of glycogen metabolizing enzymes involved in the classical GlgC-pathway and in the recently described GlgE-pathway was carried out over 47 chlamydial species representing the diversity of chlamydiae phylum. As expected, we found that a complete GlgC-pathway in most chlamydial families and the most astonishing finding was the occurrence of GlgE-pathway in three phylogenetically related Parachlamydiaceae, Waddliaceae and Criblamydiaceae families. Our genomic analysis also pinpointed a systematic lack of glgC gene in 6 draft genomes of candidatus Enkichlamydia sp. Those genomes are derived from metagenomic studies and were estimated to be 71 to 97% complete. If we assume the loss of glgC, the characterization of glycogen synthase with respect to nucleotide sugars should shed light on the glycogen pathway, as a result provide another example of the bypassing of GlgC activity in the classical GlgC-pathway. In addition to the occurrence of GlgE-pathway, a detailed genomic analysis of Waddliaceae and Criblamydiaceae families has revealed a large rearrangement of glg genes of GlgC-pathway, which had led to the loss of glgP gene and a fusion of glgA and glgB genes. This fusion appears exceptional in all three domains of life and no other such examples have been reported. In E. lausannensis (Criblamydiaceae fam.), the fusion of glgA-glgB genes is associated with a non-sense mutation resulting in premature stop codon in the open reading frame of the GlgA domain precluding the presence of the fused branching enzyme. We have shown that the glycogen synthase domain of chimeric GlgA-GlgB of W. chondrophila was active and remained ADP-glucose dependent while its branching enzyme domain already appears to be non functional due to the presence of GlgA domain at the N-terminal extremity that prevents the branching enzyme activity. In line with these observations, at variance with Parachlamydiaceae which have only maintained the glgB gene of GlgC pathway, both Waddliaceae and Criblamydiaceae have conserved glgB2 gene in GlgE operon, thereby, further suggests that the glycogen branching activity domain is indeed defective or impaired in all GlgA-GlgB fusions. Overall, this study clearly implies that GlgC-pathway does not operate in both the Waddliaceae and Criblamydiaceae families. In addition it appears possible that the genes required for the presence of a functional GlgC-pathway are at different stages of disappearance from these genomes, as suggested by the non-sense mutation in glgA-glgB gene of E. lausannensis [55].
We further investigated the GlgE glycogen biosynthesis pathway in Chlamydiales. A series of biochemical characterizations have shown that GlgE activities are capable of transferring maltosyl residue of maltose-1-phosphate onto linear chain of glucose. More remarkably, GlgE activities fulfill the priming function of glycogen biosynthesis as described for GlgA activity in GlgC-pathway [56]. We have shown that the GlgE activities switch between the processive or distributive modes of polymerization depending on the initial presence of glucan chains. Thus the “processive mode” of GlgE activity yields long glucan chains (DP>32) and is favored in their absence or in the presence of short glucan primers (DP<4). This “processive mode” of GlgE activity fills up the critical function of initiating long glucan chains that will be taken in charge by the branching enzyme in order to initiate the formation of glycogen particles. In vitro incubation experiments performed in the presence of M1P and/or branching enzyme activity further confirmed that GlgE activity is by itself sufficient for synthesizing de novo a branched polysaccharide with high molecular weight. At variance with mycobacteria and Streptomycetes, trehalose synthase (TreS) and maltokinase (Mak) activities of Chlamydiales form a bifunctional enzyme composed of TreS and Mak domains at the N- and C-terminus, respectively, which has never been reported to our knowledge. The fused TreS-Mak activity is functional and mediates the trehalose conversion into maltose and the phosphorylation of maltose into maltose-1-phosphate in the presence of ATP, GTP or UTP as phosphate donors. In contrast to mycobacteria, the maltose kinase domain requires preferentially manganese rather magnesium as divalent cation [44].
The fact that the occurrence of GlgE-pathway is limited to a few chlamydial families has led us to wonder about the origin of this operon. Our phylogeny analyses suggest that GlgE operons identified in chlamydia species share a common origin but are only distantly related to the GlgE operon from Actinobacteria (i.e Mycobacteria). We could not determine whether the presence of the GlgE pathway predated the diversification of chlamydiae or whether the operon was acquired by lateral gene transfer by the common ancestor of the Criblamydiaceae, Waddliaceae and Parachlamydiaceae from another member of PVC superphylum. One fair inference is that the genome of common ancestor of Waddliaceae, Criblamydiaceae and Parachlamydiaceae families encoded both GlgC- and GlgE-pathways. The loss of both glgP and glgC and fusion of glgA and glgB occurred before the emergence of Waddliaceae and Criblamydiaceae and may involve one single deletion event if we presume a glA/glgC/glgP/glgB gene arrangement in the common ancestor. While GlgE pathway was maintained in Waddliaceae and Criblamydiaceae due to the mandatory function of glycogen in Chlamydiales, most of members of Parachlamydiaceae retained only the GlgC-pathway except for two Protochlamydia species. The redundancy of glycogen metabolism pathway in P. naegleriophila species is quite surprising and goes against the general rule of genome optimization of intracellular obligatory bacteria. It is worthy to note that P. naegleriophila species was originally isolated from a protist Naegleria sp. N. fowleri, the etiological agent of deadly amoebic encephalitis in humans, stores carbon exclusively in the form of trehalose and is completely defective for glycogen gene network [57]. Therefore, it is tempting to hypothesize that P. neagleriophila use the retained the GlgE pathway to effectively mine the trehalose source of its host either by uptaking trehalose from its host through a putative disaccharide transporter or by secreting via type three secretion system enzymes of GlgE pathway. Our preliminary experiments based on heterologous secretion assay in Shigella flexneri suggested that GlgE and TreS-Mak could be secreted by the type three-secretion system (S8 Figure). Like Chlamydiaceae, the secretion of chlamydial glycogen metabolism pathway may be a strategy for manipulating the carbon pool of the host [54]. As reported for P. amoebophila with respect to D-glucose [49], the utpake of host’s trehalose provides an important advantage in terms of energy costs. In comparison with GlgC-pathway, one molecule of ATP is required to incorporate two glucose residues onto growing polysaccharide; at the scale of one glycogen particle synthesis this may represent a significant amount of ATP saving. The uptake of radiolabeled trehalose by host-free elementary bodies may or not support this hypothesis.
The preservation of glycogen metabolism pathway through the bottleneck of genome reduction process sheds light on a pivotal function of glycogen that has been hitherto underestimated within Chlamydiae. As a result, the question arises as to why chlamydiae have maintained glycogen metabolism pathway, making them unique among obligate intracellular bacteria. It is worthy to note that most of obligate intracellular bacteria Anaplasma spp., Ehrlichia spp., Wolbachia spp., Rickettsia spp. do not experience environmental stresses like Chlamydiae and Coxellia burnetii [58]. They thrive in nutrient rich environments either in animal or insect hosts. Losses of metabolic functions such as carbon storage metabolism in obligate intracellular bacteria are balanced by the expression of a wide variety of transporters for the uptake metabolites from the host. Except for ultra-resistant spore-like forms of C. burnetti named small cell variants, over the last decade, our perception of EB has switched from an inert spore-like form to metabolic active form capable of transcription and translation activities [49, 59]. The combination of different “omic” approaches performed on purified RB and EB of C. trachomatis and P. amoebophila have shown that genes involved in glycogen and energy metabolism pathways are upregulated in the late stage of development [4] [60][61] and most remarkably, the uptake of glucose and glucose-6-phosphate by EBs of P. amoebophila and C. trachomatis improves significantly the period of infectivity [49, 62]. Accordingly, it seems reasonable to argue that the primary function of cytosolic glycogen in EBs is to fuel metabolic processes (i.e glycolysis, pentose phosphate) when EBs are facing up the poor nutrient environment (figure 10). Future investigations should provide new opportunity to delineate the function of glycogen in chlamydiae especially with the development of forward genetic approaches [63, 64]. Finally the use of GlgE inhibitors initially designed against mycobacterial infections and to some extent the use of inhibitors of chlamydial glycogen metabolizing enzymes might define new attractive drugs to treat W. chondrophila, since this Chlamydia-related bacteria has been increasingly recognized as a human pathogen [65, 66].
Acknowledgements:
The authors are very grateful to Dr Nicolas Szydlowski for providing access to the capillary electrophoresis. We also thank Dr Agathe Subtil from Pasteur Institute (Paris) for providing Shigella strains and antibodies as well as the Plateforme d’Analyse des Glycoconjugues (PAGes, http://plateforme-pages.univ-lille1.fr/) for providing access to the instrumental facilities for carbohydrate analysis. This work was supported by the CNRS, the Université de Lille CNRS, and the ANR grants “Expendo” (ANR-14-CE11-0024).