Guanidine fuels rapid resurrection of desert cyanobacteria

Guanidine fuels rapid resurrection of desert cyanobacteria | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Biological Sciences - Article Guanidine fuels rapid resurrection of desert cyanobacteria Bao-Sheng Qiu, Jin-Long Shang, Guo-Wei Qiu, Lei Zhao, Hai-Feng Xu, and 7 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-8507520/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract In desert ecosystems, microbial activity is driven by brief hydration pulses but severely constrained by persistent nutrient scarcity 1–4 . Cyanobacteria, as essential pioneer photoautotrophs, sustain biogeochemical cycling and ecosystem stability in these arid landscapes, yet their rapid resuscitation following rewetting is critically limited by nitrogen availability 3,4 . Although the nitrogen demand is met later by biological nitrogen fixation, it is irrelevant during early rehydration due to the high energy costs and delayed nitrogenase activation, creating a critical metabolic bottleneck 5–10 . Here we demonstrate that the desert cyanobacterium Nostoc flagelliforme overcomes this constraint by activating a previously overlooked guanidine carboxylase pathway, which sustains the rapid remobilization of internal nitrogen reserves upon rehydration. Pathway activity correlates strongly with hydration and nitrogen status. Genetic experiments demonstrate its essential role in recovery from desiccation and characterize the key role of a guanidine-I riboswitch. Phylogenetic evidence indicates that pathway genes are transferred horizontally, correlating with habitat aridity. Our findings establish a novel ecological role for guanidine in desert ecosystems, and suggest its impact on microbial community assembly in nutrient-poor drylands. This work reveals a conserved adaptation mechanism that supports cyanobacterial resilience in xeric environments and provides a physiological basis for enhancing biocrust restoration strategies. Biological sciences/Microbiology/Environmental microbiology/Soil microbiology Earth and environmental sciences/Ecology/Ecophysiology cyanobacteria desert desiccation tolerance Guanidine guanidine-I riboswitch resurrection Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Introduction Desert ecosystems are characterized by extreme aridity, where microbial activity is largely restricted to brief and unpredictable hydration pulses 1–3 . The rapid resuscitation of soil microbial communities following rewetting is crucial for ecosystem functioning, yet this process is consistently limited by nitrogen (N) availability 3,4 . Although biological nitrogen fixation represents the primary nitrogen source in these environments 5 , the regulatory mechanisms governing nitrogen cycling and microbial resuscitation dynamics remain poorly understood. Cyanobacteria, as pioneer organisms in these habitats, face a critical metabolic challenge 6,7 . They must rapidly synthesize nitrogen-intensive cellular components during brief periods of moisture and light availability 2,8 . However, biological nitrogen fixation is inefficient in this initial phase due to its high energy demands and the delayed activation of the nitrogenase complex 9,10 . This contradiction between an immediate high nitrogen demand and a limited supply suggests that the efficient remobilization of internal nitrogen reserves is essential for survival. Several molecules are involved in nitrogen storage and metabolism in cyanobacteria, with cyanophycin serving as a major intracellular nitrogen reservoir 11,12 . This non-ribosomally synthesized copolymer is broken down into aspartate and arginine under nitrogen starvation 11,12 . ArgZ (AgrE), a bifunctional enzyme, then metabolizes arginine via its arginine dihydrolase activity, converting it to ornithine while releasing CO 2 and ammonia 13,14 . The resulting ornithine can either enter nitrogen assimilation via the ornithine–ammonia cycle or be converted to proline by the ornithine cyclodeaminase activity of AgrE, followed by eventual degradation via the proline oxidase PutA 13,15 . Through the combined AgrE/PutA pathway, the four nitrogen atoms of arginine are converted into three molecules of ammonia and one molecule of glutamate, thereby efficiently remobilizing nitrogen for metabolic use 15 . Recent studies have significantly advanced our understanding of nitrogen metabolism, demonstrating that some microorganisms can utilize guanidine and its derivatives as nitrogen sources 16–23 . Guanidine, a chemically stable compound containing 71.1% nitrogen by mass, was discovered more than 150 years ago 24 . However, the pathways and mechanisms underlying its biodegradation remain poorly understood. It is now known that microbes degrade guanidine primarily via two routes: the guanidinase pathway and the carboxylase pathway 17,20,21 . Both pathways ultimately convert one molecule of guanidine into three molecules of ammonia. While certain cyanobacteria and complete ammonia oxidizers (comammox) have been shown to utilize guanidine as the sole nitrogen source through the guanidinase pathway 20,21 , the physiological role of the guanidine carboxylase pathway remains unclear in cyanobacteria, and its ecological significance warrants further investigation. Here, we examined nitrogen remobilization mediated by the guanidine carboxylase pathway in the representative desert cyanobacterium Nostoc flagelliforme (hereafter N. flagelliforme ). Our results revealed that this pathway is regulated by a guanidine-I riboswitch in N. flagelliforme , a mechanism likely conserved across many other terrestrial cyanobacteria. The presence of this riboswitch-controlled system may enable desert cyanobacteria to rapidly mobilize internal nitrogen reserves during rehydration, thereby facilitating their post-desiccation recovery. This capability has important implications for biogeochemical cycling and ecosystem stability in arid and semi-arid regions. Results A guanidine-binding riboswitch is located within the 5′ UTR of the cgdA-cgdB-gcA-atzF cluster in N. flagelliforme . N. flagelliforme is a model desert cyanobacterium renowned for its extreme desiccation tolerance and rapid recovery upon rehydration 10,25 . The cgdA-cgdB-gcA-atzF gene cluster, implicated in urea degradation (Fig. 1a), was previously identified from transcriptomic data as being differentially expressed during rehydration 10 . RT‑qPCR analysis confirmed that the transcription of cgdA , cgdB , gcA , and atzF was significantly upregulated in the early phase of rehydration (Supplementary Fig. 1a). A similar induction pattern was also observed in two dominant biological soil crust cyanobacteria, Microcoleus vaginatus ( M. vaginatus , Supplementary Fig. 1b) 26 and Leptolyngbya ohadii ( L. ohadii , Supplementary Fig. 1c) 1 , suggesting a conserved role of this gene cluster during rehydration across desiccation-tolerant cyanobacteria. Bioinformatic analysis identified a putative guanidine riboswitch within the 5′ untranslated region (UTR) of this gene cluster (Fig. 1b). Secondary structure modeling revealed that it adopts the characteristic guanidine-I architecture, featuring two stem loops (P1 and P2) and a long 3′ tail overlapping a terminator hairpin (P3, Fig. 1b), consistent with known riboswitches in Sulfobacillus acidophilus 27 and Dickeya dadantii 28 . To assess the substrate-binding ability of this RNA aptamer, isothermal titration calorimetry (ITC) was performed using guanidine hydrochloride as the titrant (Fig. 1d-g). The data revealed that the 5′ UTR of N. flagelliforme cgdA binds guanidine with high affinity (Fig. 1d), but not urea (Supplementary Fig. 2). The dissociation constant ( K d ) for guanidine was determined to be 26 µM (Fig. 1d), which is lower than the previously reported value of 39.2 ± 0.85 μM in D. dadantii 28 . To exclude nonspecific binding of the RNA aptamer to guanidine, we introduced mutations into the P1 domain of the riboswitch (Fig. 1c). Deletion of the predicted guanidine-Ⅰ riboswitch (M1) abolished guanidine binding in the ITC assays (Fig. 1e), as did the disruption of the conserved base-pairing in mutant M2 (Fig. 1f). In contrast, binding was restored with the compensatory mutation M3 (Fig. 1g). Together, these results demonstrate that the 5′ UTR of the cgdA gene contains a functional riboswitch that specifically recognizes guanidine. The guanidine riboswitch regulates expression of the cgdA-cgdB-gcA-atzF gene cluster. Bioinformatic prediction of an NtcA binding site (5′-GTAgtaagtgaTAC-3′) within the cgdA promoter region (Supplementary Fig. 3) indicated that the expression of cgdA in N. flagelliforme is likely regulated by the global nitrogen regulator NtcA 29–31 . This interaction was validated by an electrophoretic mobility shift assay (EMSA; Supplementary Fig. 3c). To investigate the function of the guanidine riboswitch in vivo , site-directed mutations were introduced into its native locus using CRISPR-Cpf1 genome editing 32 (Fig. 2a-c). RT-qPCR analysis revealed that in wild-type background controls ( cgdA:: Ctrl), genes downstream of the promoter were significantly induced under nitrate starvation (Fig. 2a), consistent with the typical expression pattern of the NtcA regulon. Notably, this induction was abolished in the cgdA:: M2 mutant (Fig. 2b). The loss of induction was specifically attributable to the disruption of the riboswitch and not to any other effects, as evidenced by the restored gene induction in the compensatory mutant cgdA ::M3 (Fig. 2c). The transcriptional response of the cgdA-cgdB-gcA-atzF gene cluster was further assessed using a luciferase reporter system. In this assay, promoter activity was measured by quantifying LuxAB-derived bioluminescence following fusion to different promoter variants, each of which retained the NtcA-binding site. Compared to cultures grown under nitrogen-replete conditions (BG11, day 0), reporter strains carrying the wild-type cgdA promoter exhibited a rapid increase in LuxAB bioluminescence during the first three days of incubation in nitrate-depleted medium (BG11 0 ; Fig. 2d). Although the NtcA-binding site remained intact, no increase in bioluminescence was observed in strains with an inactivated guanidine riboswitch (M2; Fig. 2e). In contrast, introducing a compensatory mutation (M3) restored nitrogen-starvation-induced expression of cgdA (Fig. 2f). Together, these findings underscore the physiological relevance of the cgdA-cgdB-gcA-atzF gene cluster in the low-N response, as well as the essential role of the guanidine riboswitch in regulating it. The expression of the cgdA-cgdB-gcA-atzF cluster is also regulated by dehydration and rehydration. The transcriptional dynamics of the cgdA - cgdB - gcA - atzF gene cluster during dehydration and rehydration cycles were further examined using the same luciferase reporter strains (Fig. 3a-f). In this assay, N. flagelliforme cells were first cultured for 3 days in liquid BG11 or BG11 0 medium, collected on a nitrocellulose membrane, and subjected to periodic dehydration and rehydration cycles (21 h: 3 h). Dehydration was achieved by air-drying membranes at 70% relative humidity, while rehydration was performed by placing membranes on 1.5% (m/v) BG11 agar plates with or without nitrate 33 . As anticipated, nitrogen-starved reporter strains carrying either the wild-type (Fig. 3a) or the compensatory M3 (Fig. 3c) riboswitch exhibited stronger luminescence during the dehydration and rehydration cycles. In contrast, the luminescence signal from the nitrogen-depleted M2- luxAB strain remained similar to that observed under nitrogen-replete conditions (Fig. 3b). To comprehensively assess cgdA promoter activity under fluctuating water availability, time-course bioluminescence analysis was performed on day 6 (the third dehydration-rehydration cycle). As shown in Fig. 3d, bioluminescence increased during rehydration and decreased during dehydration. Notably, this oscillatory expression pattern was not of circadian origin. When rehydration was extended by an additional three hours, cgdA promoter activity remained elevated and only decreased upon re-exposure to dehydration (Supplementary Fig. 4). Consistently, bioluminescence induction was absent in the guanidine riboswitch-inactivated strain (M2; Fig. 3e) but was restored in the compensatory mutant (M3; Fig. 3f). These results indicate that the guanidine riboswitch is crucial for activating the cgdA - cgdB - gcA - atzF gene cluster during rehydration of nitrogen-starved N. flagelliforme . The cgdA-cgdB-gcA-atzF cluster is responsible for guanidine carboxylation in N. flagelliforme . While genome annotations have linked the cgdA-cgdB-gcA-atzF cluster to the urea carboxylation process, a recent study in gamma proteobacteria suggested its potential involvement in guanidine carboxylation 17 . Consistent with this, our ITC data demonstrated that the 5′ UTR of this cluster specifically binds guanidine but not urea, indicating its role in guanidine metabolism. To test whether a functional guanidine carboxylation pathway exists in N. flagelliforme , we employed U-[ 13 C, 15 N]-arginine as a tracer. In the presence of ethylene-forming enzyme (Efe) performing a reaction as described for the Pseudomonas savastanoi Efe (Supplementary Fig. 5) 34 , arginine should be converted to guanidine labeled accordingly (Fig. 4). LC-MS analysis successfully detected 15 N- and 13 C-labeled guanidine, as well as its isotopically labeled degradation products, carboxyguanidine and allophanate (Fig. 4), providing direct evidence for the presence of an arginine-to-guanidine Efe activity and of an active guanidine carboxylation pathway in N. flagelliforme . We also compared the metabolism of the wild-type ( cgdA ::Ctrl), riboswitch mutant ( cgdA ::M2), and complementary strain ( cgdA ::M3) during the rewetting phase following cycles of dehydration and rehydration. After three days of periodic dehydration and rehydration, the cgdA ::M2 mutant accumulated significantly less intracellular arginine (Fig. 3g). Interestingly, arginine levels in this mutant increased upon rehydration (Fig. 3g), likely due to nitrogen remobilization via the cyanophycin breakdown pathway. In contrast, such arginine accumulation was not observed in the cgdA ::Ctrl and cgdA ::M3 strains (Fig. 3g), suggesting that in these strains, arginine was rapidly converted into guanidine and succinate by the guanidine biosynthesis protein Efe. This was supported by measurements of guanidine and succinate levels (Fig. 3 h-i). Because the cgdA ::M2 strain could not induce guanidine carboxylation genes during rehydration, it accumulated substantially more guanidine than the cgdA ::Ctrl and cgdA ::M3 strains (Fig. 3h). Together, these results indicate that the cgdA-cgdB-gcA-atzF cluster is essential for guanidine breakdown by N. flagelliforme . Guanidine carboxylation plays an essential role in adaptation to dehydration and rehydration cycles. To investigate the physiological role of the guanidine carboxylation pathway, we measured the photosynthetic performance of N. flagelliforme strains ( cgdA ::Ctrl, cgdA ::M2, and cgdA ::M3) after 3 hours of recovery following successive dehydration and rehydration cycles. After three days of treatment, the cgdA:: M2 mutant exhibited significantly lower rates of oxygen evolution and respiration (Supplementary Fig. 6a). The phycobilisome absorbance in the cgdA:: M2 strain was also lower than that in the cgdA ::Ctrl and cgdA ::M3 strains (Supplementary Fig. 6b). Additionally, nitrogenase activity in cgdA ::M2 was lower than that in the other two strains (Supplementary Fig. 6c). These diminished metabolic activities likely resulted from the impaired recovery of the cgdA:: M2 strain during rehydration, as time-series chlorophyll fluorescence measurements showed that this riboswitch-inactivated mutant exhibited slower photosynthetic recovery, especially after multiple rounds of dehydration and rehydration (Supplementary Fig. 6d). To further understand the transcriptional responses during recovery, we analyzed the transcriptomic profiles of N. flagelliforme following successive dehydration–rehydration cycles. Based on Fuzzy C-Means clustering (FCM; Supplementary Fig. 7), genes associated with ribosomal function, DNA replication, recombination, and repair were upregulated in the cgdA ::M2 strain during rehydration (Cluster 2; Supplementary Table 1). In contrast, genes involved in photosynthesis, nitrogen fixation, hydrogenase activity, and the guanidine carboxylation pathway were downregulated (Cluster 4; Supplementary Table 1 and Supplementary Figs. 7-9), consistent with the physiological differences observed between cgdA ::Ctrl and cgdA ::M2 strains (Supplementary Fig. 6). Further analysis showed that the nitrogen‑limitation biomarker genes nrtA (FC=1.56, P <0.05), nrtB (FC=2.33, P <0.05), and amt1 (FC=1.73, P <0.05) were more highly expressed in cgdA ::M2 (Supplementary Data 1), indicating a more constrained nitrogen supply in the riboswitch‑inactivated mutant. Consistent with this, the quantification of intracellular cyanophycin revealed that N. flagelliforme strains unable to induce guanidine carboxylase (M2) exhibited reduced cyanophycin degradation efficiency (Supplementary Fig. 10). Together, these results demonstrate that disruption of the guanidine carboxylation pathway significantly impairs nitrogen remobilization during rehydration. After six days of periodic dehydration and rehydration, cgdA ::M2 strains grown under nitrogen-replete conditions exhibited a growth pattern similar to that of cgdA ::Ctrl and cgdA ::M3 strains (Fig. 5a). In contrast, under nitrogen-deprived conditions, the cgdA ::M2 strain accumulated less biomass and displayed more pronounced chlorosis than the other two strains following the same treatment (Fig. 5b), despite the higher expression of argZ in the mutant (Supplementary Data 1). When recovered in liquid BG11 medium (Fig. 5c-f), only the nitrogen-starved cgdA ::Ctrl and cgdA ::M3 strains were able to rapidly resume growth and regain green pigmentation after six days, whereas the resuscitation rate of nitrogen-starved cgdA ::M2 was markedly lower (Fig. 5d, f). These results indicate that the riboswitch-regulated guanidine carboxylation pathway is essential for cell survival and recovery during dehydration and rehydration cycles. The guanidine carboxylation gene cluster originated from an ancient horizontal gene transfer event. To elucidate the evolutionary origin of the cyanobacterial guanidine carboxylation pathway, we assembled a comprehensive dataset comprising 16S rDNA sequences and concatenated CgdA and CgdB protein sequences. Phylogenetic reconstruction revealed that the maximum-likelihood species tree was largely congruent with the guanidine catabolism tree inferred from the concatenated CgdA and CgdB alignment (Supplementary Figs. 11, 12). All cyanobacterial guanidine carboxylation genes formed a distinct monophyletic clade separate from those of other bacteria (Supplementary Fig. 12), implying that cyanobacteria likely acquired an ancestral carboxylation module for guanidine degradation from proteobacteria via horizontal gene transfer (HGT) 10,35 , followed by gene duplication and functional diversification into different cyanobacterial lineages. Moreover, phylogenetic comparisons indicate that the marker genes have coevolved with the guanidine‑degradation genes in cyanobacteria, as supported by their high topological congruence (Supplementary Fig. 13). Together, these results suggest that the HGT‑mediated acquisition of an ancestral carboxylation reaction, which initiates guanidine degradation, likely facilitated the adaptive radiation of these cyanobacteria into dynamic habitats. The riboswitch-regulated guanidine carboxylase pathway is predominantly found in terrestrial cyanobacteria. To examine the distribution of the riboswitch-regulated guanidine carboxylase pathway in cyanobacteria, we performed comparative genomic analyses of strains from diverse representative habitats. Homology searches across 2020 cyanobacterial strains identified 835 unique guanidine-I RNA sequence motifs (RF00442 in the Rfam database) distributed among 739 strains (Supplementary Table 2 and Supplementary Fig. 14). While most of these cyanobacteria contain a single copy of the guanidine-I riboswitch, some strains carry two or three distinct copies (Fig. 6 and Supplementary Table 2). These results indicate that the guanidine-I riboswitches are widely distributed across the cyanobacterial phylum. To identify genes linked to the guanidine‑I aptamer, we compared the sequences immediately downstream of the aptamer with the NCBI protein database. Among the 739 genomes analyzed, the aptamer was located upstream of gdmH (encoding guanidine hydrolase 20 ) in 648 cases and upstream of cgdAB (encoding carboxyguanidine deiminase) in 91 cases (Supplementary Table 2). Intriguingly, although both pathways are involved in guanidine utilization, they appear to be mutually exclusive, as no co-occurrence of the two has been observed. Further analysis revealed that the guanidine‑I riboswitch-regulated carboxylase pathway was predominantly distributed in terrestrial cyanobacteria (Fig. 6c and Supplementary Table 2). Moreover, its presence shows a positive association with habitat aridity (Fig. 6c). Discussion Desert ecosystems exhibit low nitrogen bioavailability, due to limited external inputs and incomplete nitrogen cycling 36–38 . This nitrogen limitation constrains microbial activity, shapes the soil community structure, and ultimately restricts primary productivity. To survive such extremes, desert cyanobacteria must rapidly remobilize intracellular nitrogen when water becomes available. However, the mechanism by which they acquire usable nitrogen during rehydration has remained enigmatic. Here, we present multiple evidence that the riboswitch-controlled guanidine carboxylase pathway is essential for nitrogen remobilization upon rehydration. Although the hydrolysis pathway appears to be more common in aquatic cyanobacteria, the carboxylase pathway is ecologically significant in terrestrial habitats. This efficient nitrogen-remobilization pathway likely enables desert cyanobacteria to swiftly restore cellular functions during the brief periods when both water and light are available, thus priming them for subsequent dehydration. Several metabolic pathways have been implicated in the mobilization of intracellular nitrogen reserves in cyanobacteria. Transcriptomic analysis revealed high expression of agrE ( argZ ) and putA in both cgdA:: Ctrl and cgdA:: M2 strains (Supplementary Data 1), underscoring their importance in nitrogen remobilization during the rehydration process. Although the AgrE/PutA pathway is highly efficient 15 , our data show that N. flagelliforme strains unable to upregulate the guanidine carboxylase pathway ( cgdA ::M2) displayed significantly reduced cellular activity and slower photosynthetic recovery, even though agrE expression was higher than in the wild‑type (Supplementary Fig. 9). These results indicate that arginine catabolism via the AgrE/PutA pathway alone cannot meet the high nitrogen demand of rehydrating cyanobacteria, and further highlight the guanidine carboxylase pathway as a critical mechanism for supporting elevated nitrogen requirements and fueling cellular recovery. Moreover, in contrast to arginine remobilization via the AgrE/PutA pathway and the arginine‑ammonia cycle 13,15 , the Efe-catalyzed conversion of arginine to guanidine consumes three molecules of α-ketoglutarate (2-OG; Fig. 4a) 34 . Given the central role of 2‑OG in regulating carbon-nitrogen metabolic balance and its marked accumulation under nitrogen-limited conditions 39–42 , guanidine-dependent arginine degradation may provide a route for nitrogen-starved cyanobacteria to simultaneously catabolize stored carbon and rapidly restore metabolic homeostasis. While the role of NtcA in nitrogen homeostasis is widely recognized 29–31 , our findings elucidate an additional layer of regulation mediated by the guanidine‑I riboswitch for guanidine carboxylation and degradation. ITC analysis confirmed that the guanidine‑I riboswitch in the N. flagelliforme carboxylase gene cluster specifically binds guanidine, but not urea, validating the precision of this metabolite‑dependent switch. Notably, homologous riboswitches are commonly found upstream of guanidine carboxylase or hydrolase clusters in diverse cyanobacterial strains (Supplementary Table 2). This consistent genomic association suggests that the riboswitch‑dependent regulatory mechanism described here is likely conserved across cyanobacteria. Interestingly, analysis of guanidine carboxylase gene distribution shows that guanidine‑I riboswitch‑controlled carboxylase pathways are primarily found in terrestrial cyanobacteria (Fig. 6c). This pattern matches the ecological niche of Pseudomonas syringae 17 , the model organism for guanidine carboxylation, but differs markedly from that of nickel (Ni)-dependent guanidine hydrolase genes, which are more frequently found in aquatic environments (Fig. 6c). Although Ni is generally scarce in both terrestrial and aquatic environments, its concentration in the ocean exceeds that of other essential trace metals, such as Cu, Fe, and Mn 43,44 . In marine systems, low Ni availability does not appear to drive the loss of Ni-dependent pathways. Instead, marine cyanobacteria preferentially utilize Ni-dependent superoxide dismutase (NiSOD) over SODs that rely on other metal cofactors 45,46 . In contrast, many terrestrial ecosystems exhibit particularly low Ni bioavailability 47,48 . In alkaline soils of arid and semi-arid regions, where precipitation-driven acidification and leaching are minimal, Ni deficiency is likely widespread. Under these conditions, replacing Ni-dependent guanidine hydrolase with a carboxylase pathway may have relieved desert cyanobacteria of the metabolic burden of acquiring scarce nickel 17,20,21 . Moreover, the Michaelis constant ( K m ) of Pseudomonas syringae GcA for guanidine is 0.21 ± 0.09 mM 17 , which is approximately an order of magnitude lower than the values reported for guanidinase in Synechocystis sp. PCC 6803 (7.81 ± 1.80 mM) 20 and Nitrospira inopinata (13.6 ± 0.76 mM) 21 . This lower K m of the carboxylase at physiological pH reflects a high catalytic efficiency, enabling effective guanidine utilization even at low substrate concentrations during rehydration. Recent studies have significantly advanced our understanding of guanidine biosynthesis 49 . In addition to the reaction catalyzed by Efe, microorganisms are known to produce guanidine during naphthyridinomycin biosynthesis and through the complete degradation of metformin, one of the world’s most widely prescribed drugs 23,50–52 . In land plants, homoarginine‑6‑hydroxylases are estimated to generate approximately 3.5 × 10 5 tons of guanidine annually 53 . However, plants appear to primarily transport rather than metabolize guanidine 54 , suggesting that its accumulation or secretion may serve as a strategy to modulate associated microbial communities 55 . Intriguingly, our analysis indicated that some terrestrial cyanobacteria lacking known guanidine‑producing genes harbor gene clusters encoding ATP‑binding cassette (ABC)‑type guanidine transporters (Fig. 6). For instance, Microcoleus vaginatus lacks identified guanidine synthesis genes, yet it encodes both the guanidine carboxylation pathway and a high‑affinity guanidine importer, and harbors a guanidine‑I riboswitch upstream of its ABC transporter locus (Supplementary Fig. 15). Microcoleus species are representative biocrust cyanobacteria that do not fix dinitrogen yet dominate nitrogen‑limited biological soil crusts 26 . Although nutrient exchange with associated microbes is well documented in Microcoleus and urea has been proposed as the main vehicle for interspecies nitrogen transfer 56–59 , our analysis suggests that its annotated urea carboxylase is more likely to function in guanidine carboxylation (Supplementary Fig. 15c). Thus, guanidine-based mutualistic nitrogen transfer within biocrusts merits further investigation. Collectively, our findings shed light on the physiological role of guanidine catabolism during rehydration. The guanidine‑I riboswitch controlled carboxylation pathway exhibits high nitrogen remobilization efficiency and restricted phylogenetic distribution outside bacteria 20 . This indicates that it gives N. flagelliforme a competitive advantage in nitrogen-poor deserts. This metabolic adaptation could be critical for cyanobacterial survival in these environments, with broader implications for primary productivity and biogeochemical cycling in dryland ecosystems. Materials and methods Cyanobacterial strains and culture conditions. The axenic strain Nostoc flagelliforme CCNUN1 was isolated from dried field samples collected in Sunitezuoqi, Inner Mongolia, China 60 . All strains used in this study are listed in Supplementary Table 5. Unless otherwise specified, cultures were grown in BG11 medium at 25°C under continuous cool-white fluorescent light (15 μmol photons m −2 s −1 ). Mutants of N. flagelliforme were maintained in BG11 medium supplemented with 50 μg mL −1 spectinomycin, and no antibiotics were added during the experimental treatments. To simulate the natural dehydration-rehydration cycle of N. flagelliforme , exponentially growing cultures pre-adapted to nitrogen-replete or nitrogen-deplete medium were collected on 0.45 μm nitrocellulose membrane filters (Millipore), respectively. The samples were air-dried for 21 hours at 25°C, 70% relative humidity, and an irradiance of 15 μmol photons m −2 s −1 , followed by rehydration on BG11 or BG11 0 (combined nitrogen-free) plates containing 1.5% agarose for 3 hours. This drying-rewetting cycle was repeated six times. Subsequently, the strains were recovered in liquid BG11 medium for assessment of cellular activity. RNA extraction and RT-qPCR analysis. Total RNA was extracted from N. flagelliforme using Trizol Reagent (Cwbio, China) 61 . Genomic DNA was digested, and cDNA was synthesized using the PrimeScript reagent gDNA Eraser Kit (Takara) according to the manufacturer’s instructions. Transcript levels of target genes were quantified by RT-qPCR using a 7900HT Fast Real-Time PCR System (Thermo Fisher Scientific, USA) with SYBR Green Real-Time PCR Master Mix (Toyobo). The 16S rDNA gene was used as an internal reference gene. Relative expression levels were calculated using the 2 −ΔΔCT method. All primers used are listed in Supplementary Table 4. RNA preparation and ITC analysis. The DNA fragment containing the 5′ UTR of cgdA (designated Ctrl) was amplified using specific primers. Corresponding mutations (M1, M2, and M3) were then introduced into this fragment via fusion PCR. For in vitro transcription, 20 μL reactions were prepared containing 50 mM Tris-HCl (pH 7.5), 15 mM MgCl 2 , 2 mM spermidine, 5 mM DTT, 2.5 mM of each ribonucleoside 5′ triphosphates (NTP), and 20 units of T7 RNA polymerase (Vazyme Biotech) 62 . Reactions were incubated at 37°C for 2 h, and the resulting RNA was purified using the RNA Isolation Kit V2 (Vazyme Biotech) according to the manufacturer’s instructions. Isothermal titration calorimetry (ITC) experiments were conducted using a TA Instruments Nano ITC equipped with a low-volume sample cell at 25°C 28 . Guanidine hydrochloride (Sigma-Aldrich) was dissolved in Milli‑Q water (18.2 MΩ cm −1 ) to a concentration of 1.4 mM and then titrated into the sample cell containing 0.2 mM RNA. A total of 25 serial injections of 2 μL each were performed, with 180‑second intervals between injections. Data were fitted to a one‑site binding model and analyzed using the Nanoanalyze software. Luciferase reporter assays. The luciferase reporter system was constructed as described below. Using the Ctrl- luxAB strain as an example, the 5′ UTR of cgdA was fused to the luxAB reporter fragment. The resulting DNA fragment was cloned into the pRL25C vector to generate the recombinant plasmid pRL25C-Omega- Promoter - luxAB . This plasmid was introduced into N. flagelliforme CCNUN1 using the conjugation method 63 . Transformants were selected on 1.5% agar BG11 plates containing 50 μg mL −1 spectinomycin under 15 μmol photons m −2 s −1 illumination. Positive single colonies were verified by PCR and sequencing. Primers used are listed in Supplementary Table 4, and the resulting plasmids and strains are summarized in Supplementary Table 5. Promoter activity was determined by measuring the relative bioluminescence intensity of luciferase in N. flagelliforme CCNUN1 cells 61 . Aliquots of 0.2 mL culture, each containing an equal amount of chlorophyll a , were transferred to a 96-well plate. Then, 5 μL of 0.1 mM decanal (dissolved in dimethyl sulphoxide) was added as the substrate. Bioluminescence was measured at room temperature using a BioTek Synergy 2 Plate Reader (BioTek). The recorded luminescence values were normalized to the corresponding chlorophyll a content. Mutant construction. Chromosomal mutagenesis of the guanidine riboswitch in N. flagelliforme was performed using CRISPR-Cpf1 genome editing 32 . Briefly, the “TTN” motif within the guanidine‑I aptamer was identified, and a 20 bp DNA sequence downstream of this motif was inserted into the AarⅠ site of the pCpf1 vector. Subsequently, the wild‑type fragment (Ctrl) and the mutated fragments (M2 and M3) were separately cloned into the BglⅡ site of the same vector. The resulting plasmids were transformed into E. coli HB101 and then transferred into N. flagelliforme via conjugation 63 . Mutants were selected on BG11 agar plates containing 50 μg mL −1 spectinomycin under 15 µmol photons m −2 s −1 . Positive recombinant strains were verified using PCR and sequencing. All primers are listed in Supplementary Table 4, and the resulting plasmids and strains are summarized in Supplementary Table 5. Metabolic analysis and isotope labeling experiments. The N. flagelliforme strains that had undergone periodic dehydration and rehydration were scraped from the filter membrane and quenched in cold methanol 64 . After centrifugation, the pellet was ground in liquid nitrogen, followed by three rounds of extraction with 500 μL of a 50:50 (v/v) methanol:water mixture. The supernatants from each extraction were pooled, dried in a vacuum centrifuge, and reconstituted in 500 μL Milli-Q water (18.2 MΩ cm −1 ). The resulting extracts were filtered through a 0.22 μm nylon membrane filter (ANPEL, Shanghai, China) prior to analysis. Metabolite extracts were analyzed using a Shimadzu UFLC-30A High-Performance Liquid Chromatograph (HPLC) system coupled to an Applied Biosystems 4500 QTRAP mass spectrometer (MS). Chromatographic separation was performed using a Waters ACQUITY UPLC® BEH Amide column (2.1×100 mm, 1.7 µm particle size). The mobile phase consisted of solvent A (water containing 10% ammonium hydroxide and 50 mM ammonium formate) and solvent B (acetonitrile). A linear gradient was applied as follows: solvent A was initially set at 10%, increased to 90% over 9 min, held for 1 min, returned to 10% within 0.5 min, and maintained for 2.5 min. The injection volume was 4 µL, flow rate was 0.4 mL min −1 , and column temperature was maintained at 40°C. Metabolites were detected in the positive ion mode using a quadrupole-linear ion trap mass spectrometer. Data were acquired using the AB Sciex Analyst 1.6 software. To examine the potential presence of a guanidine carboxylation pathway, N. flagelliforme was first pre‑adapted in nitrogen‑deplete medium and harvested onto 0.45 μm nitrocellulose membrane filters. The samples were air‑dried for dehydration and then rehydrated on a BG11 0 plate supplemented with 1 mM uniformly labeled U-[ 13 C, 15 N]-arginine. Following 3 days of repeated dehydration–rehydration cycles, the samples were collected for analysis. U-[ 13 C, 15 N]-arginine was purchased from Cambridge Isotope Lab, and guanidine hydrochloride, arginine, and succinate were obtained from Sigma‑Aldrich. Standard solutions were prepared at defined concentrations, and a subset of the identified compounds was verified by matching their mass and retention time to those of authenticated standards. Determination of photosynthetic activity. Photosynthetic oxygen evolution was measured using a Clark-type oxygen electrode (Chlorolab 2, Hansatech Instruments) 33 . After periodic dehydration and rehydration, samples were scraped from the filters and resuspended in the reaction medium (BG11 medium buffered with 25 mM Bis‑Tris propane, pH 8.0, and supplemented with 1 mM NaHCO 3 ). The electrode was calibrated with air‑equilibrated distilled water (maximum signal) and with a Na 2 S 2 O 4 solution (minimum signal). Temperature was maintained at 25°C using a Polystat refrigerated bath (Cole-Parmer Instrument, Vernon Hills, IL). Net O 2 evolution was measured at 960 μmol photons m −2 s −1 , and respiration was measured as O 2 consumption in the dark. Chlorophyll fluorescence was measured with a Plant Efficiency Analyser (Hansatech Instruments). Prior to measurement, samples were dark-adapted for 15 min to ensure that all PSII reaction centers were in the open state 33 . The maximal fluorescence ( F m ) was determined by applying a pulse of saturating red light, and the maximum quantum yield of PSII ( F v / F m ) was calculated as ( F m – F o )/ F m . Measurement of nitrogenase activity. Nitrogenase activity in N. flagelliforme strains was determined using the acetylene reduction assay 65 . Following treatment, cells were scraped from the filter and resuspended in 2 mL of BG11 0 medium inside a 10‑mL sealed tube and adjusted to a final chlorophyll a content of 0.1 mg. The headspace of each tube was repeatedly evacuated and replaced with C 2 H 2 gas. Samples were then incubated at 5 μmol photons m −2 s −1 and 25°C for 6 h, after which the gas phase was collected and analyzed on an Agilent 7650A gas chromatograph. Data are presented as mean values from at least three replicate cultures. Transcriptomic analysis. Triplicate cultures of cgdA:: Ctrl and cgdA:: M2 strains were collected after rehydration following dehydration and were immediately frozen in liquid nitrogen. Total RNA was extracted using TRIzol Reagent according to the manufacturer’s instructions (Invitrogen). RNA concentration was measured using an ND-2000 spectrophotometer (NanoDrop), and the samples were subsequently sent to Majorbio Co. Ltd. for library preparation and 150‑bp paired-end sequencing on an Illumina HiSeq×TEN platform. Ribosomal RNA sequences were removed using SortMeRNA v1.9 66 , and the resulting clean reads were aligned to the reference genome using Rockhopper 67 . Normalization factors were calculated using the Trimmed Mean of M‑values method, and differential expression analysis was performed with edgeR v3.20.7 68 . Gene expression profiles were further analyzed statistically using the Mfuzz package from Bioconductor in R 69 . A full list of the differentially expressed genes is available in Supplementary Data 1. In silico analysis of guanidine catabolism genes. Homologs of CgdA, CgdB, GcA, AtzF, GdmH, GhaA, GhaB, ABC1-3, SssF, and Efe were identified using BLAST searches against the NCBI databases. For phylogenetic analysis, two datasets were separately aligned using the MAFFT multiple alignment program 70 : one comprising a concatenation of 31 single‑copy protein‑coding genes conserved across all representative species, and the other consisting of 16S rDNA sequences from representative cyanobacteria and bacteria. A maximum‑likelihood phylogenetic tree was constructed with RAxML v8.1.20 under the PROTGAMMA model, with branch support evaluated by 1,000 bootstrap replicates, as previously described 10 . Basic genomic statistics for the 2,020 cyanobacterial genomes used in the comparative analysis are provided in Supplementary Table 2. The occurrence and frequency of the RNA motif RF00442 (guanidine‑I aptamer) were examined using Infernal (v.1.1.2), with CMsearch applied to the motif at an E -value threshold of 0.0001 against all replicons from the included species 71 . Statistical analyses. All experiments were performed with a minimum of three independent replicates. Statistical processing was conducted using Origin 2021 software (OriginLab). For comparisons across multiple treatment groups, Tukey’s honest significant difference (HSD) test was applied, with a P -value<0.05 considered statistically significant. Differences between two independent groups were evaluated using the Student’s t -test. Declarations Data availability. Data supporting the findings of this study are provided in the Supplementary Data files. Transcriptomic data have been deposited in the NCBI SRA database under accession code PRJNA1099419: https://www.ncbi.nlm.nih.gov/search/all/?term=PRJNA1099419. Guanidine-I aptamers and guanidine catabolism genes were identified using BLAST searches of NCBI databases, and the corresponding results are presented in Tables S2 and S3. Acknowledgement This work was funded by the National Natural Science Foundation of China (No. 32430005, No. 32270397, and No. 32470408), the China Postdoctoral Science Foundation (No. 2019T120675), and the Tianchi Talent Program of the Xinjiang Uygur Autonomous Region. Author contributions : J.L.S., and B.S.Q. designed the research project; J.L.S., L.Z., Y.C., and Y.L. performed research; J.L.S., G.W.Q., L.Z., H.F.X., Z.C.Z., G.Z.D., S.H., C.Y., W.R.H., and B.S.Q. analyzed data; and J.L.S., G.W.Q., W.R.H., and B.S.Q. wrote the paper. The authors declare no competing interest. References Oren, N., Raanan, H., Murik, O., Keren, N. & Kaplan, A. Dawn illumination prepares desert cyanobacteria for dehydration. Curr. Biol. 27 , R1056–R1057 (2017). Xu, H. F. et al. Reading and surviving the harsh conditions in desert biological soil crust: the cyanobacterial viewpoint. FEMS Microbiol. Rev. 45 , fuab036 (2021). Imminger, S. et al. 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MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Mol. Biol. Evol. 30 , 772–780 (2013). Nawrocki, E. P. & Eddy, S. R. Infernal 1.1: 100-fold faster RNA homology searches. Bioinformatics 29 , 2933–2935 (2013). Additional Declarations There is NO Competing Interest. Supplementary Files SupplementaryData1.xlsx Supplementary Data SupplementaryFigures.pdf Supplementary Figures 1-15 SupplementaryTableS1XXXS5.xlsx Supplementary Tables S1–S5 Cite Share Download PDF Status: Posted Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. 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UTR of the \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003ecgdA\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e gene in\u003c/strong\u003e\u003cem\u003e\u003cstrong\u003e N. flagelliforme\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e contains a guanidine-binding aptamer. a\u003c/strong\u003e The schematic diagram illustrates the genomic organization of the genes encoding CgdA, CgdB, GcA, and AtzF in the genome of \u003cem\u003eN. flagelliforme\u003c/em\u003e. A putative open reading frame of unknown function is shown in light gray, and the gene encoding an amidase is marked in brown. The arrow indicates the predicted NtcA-binding site. \u003cstrong\u003eb\u003c/strong\u003e Predicted secondary RNA structure of the guanidine-I riboswitch located in the 5′UTR of the \u003cem\u003ecgdA\u003c/em\u003e gene in\u003cem\u003e N. flagelliforme\u003c/em\u003e. \u003cstrong\u003ec\u003c/strong\u003e Schematic representation of the riboswitch mutagenesis strategies. In the M1 mutant, the entire guanidine-I riboswitch was deleted from the 5′ UTR of the \u003cem\u003ecgdA\u003c/em\u003e gene. In mutants M2 and M3, site-directed mutations were introduced into conserved nucleotides within the P1 stem, with M3 containing compensatory mutations to restore base pairing. \u003cstrong\u003ed\u003c/strong\u003e-\u003cstrong\u003eg\u003c/strong\u003e Binding affinities of the wild-type \u003cem\u003ecdgA\u003c/em\u003e riboswitch (Ctrl, \u003cstrong\u003ed\u003c/strong\u003e) and its mutants (M1-M3, \u003cstrong\u003ee\u003c/strong\u003e-\u003cstrong\u003eg\u003c/strong\u003e) for guanidine. Upper panels show raw isothermal titration calorimetry data; lower panels display the integrated heat measurements. Binding isotherms and fitted curves were derived from a one-site binding model. Data are from one representative experiment out of three independent biological replicates.\u003c/p\u003e","description":"","filename":"Figures1.png","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/55185979b5adf07e441c4b35.png"},{"id":100869390,"identity":"29fdc2a2-5dfc-408d-b330-16abe87ca781","added_by":"auto","created_at":"2026-01-22 08:59:02","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":77106,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eGuanidine riboswitch mediates \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003ecgdA\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e induction under nitrogen starvation. a\u003c/strong\u003e-\u003cstrong\u003ec\u003c/strong\u003e Transcript levels of \u003cem\u003ecgdA\u003c/em\u003e, \u003cem\u003ecgdB\u003c/em\u003e, \u003cem\u003egcA\u003c/em\u003e, and \u003cem\u003eatzF\u003c/em\u003e\u003cstrong\u003e \u003c/strong\u003ein the wild-type (\u003cem\u003ecgdA\u003c/em\u003e::Ctrl,\u003cstrong\u003e a\u003c/strong\u003e), the riboswitch mutant (\u003cem\u003ecgdA\u003c/em\u003e::M2, \u003cstrong\u003eb\u003c/strong\u003e), and the complementary strain (\u003cem\u003ecgdA\u003c/em\u003e::M3, \u003cstrong\u003ec\u003c/strong\u003e) during a 3-day incubation under nitrogen-deficient conditions. The upper panels depict the mutation strategies introduced into the \u003cem\u003eN. flagelliforme\u003c/em\u003e riboswitch, with conserved nucleotides altered in the P1 stem highlighted in red. Data are presented as mean ± SD from three independent biological replicates, normalized to transcript levels measured under nitrogen-sufficient conditions. \u003cstrong\u003ed\u003c/strong\u003e-\u003cstrong\u003ef\u003c/strong\u003e Promoter activity of the \u003cem\u003ecgdA\u003c/em\u003e gene under nitrogen deficiency. Activity was quantified by measuring bioluminescence produced by a LuxAB reporter fused to each promoter variant. Data are shown as mean ± SD of three independent biological replicates. Letters above bars denote statistically significant differences (\u003cem\u003eP\u003c/em\u003e \u0026lt; 0.05) determined by one-way ANOVA across all treatments.\u003c/p\u003e","description":"","filename":"Figures2.png","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/895b20725898b0f4cf9d8ed5.png"},{"id":102294777,"identity":"62f96055-7f27-4257-8c2f-7ea13415c5c2","added_by":"auto","created_at":"2026-02-10 09:56:26","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":68154,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eGuanidine-I riboswitch mutagenesis blocks \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003ecgdA\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e induction and consequently causes guanidine accumulation upon rehydration. a\u003c/strong\u003e-\u003cstrong\u003ef\u003c/strong\u003e Promoter activity of the \u003cem\u003ecgdA\u003c/em\u003e gene\u003cem\u003e \u003c/em\u003eunder nitrogen limitation and rehydration. \u003cem\u003eN. flagelliforme\u003c/em\u003e cells carrying a LuxAB reporter fused to different promoter variants were grown exponentially in liquid BG11 or BG11\u003csub\u003e0\u003c/sub\u003e (BG11 without nitrate) medium for 3 days. Cells were then collected on nitrocellulose membrane filters and subjected to periodic dehydration and rehydration cycles (21 h: 3 h) for an additional 5 days. Promoter activity was quantified daily as relative luminescence units (RLU) of luciferase (\u003cstrong\u003ea\u003c/strong\u003e-\u003cstrong\u003ec)\u003c/strong\u003e. A time‑course bioluminescence analysis was performed on the third day of the dehydration-rehydration cycles to assess promoter dynamics during these phases (\u003cstrong\u003ed\u003c/strong\u003e-\u003cstrong\u003ef\u003c/strong\u003e). Data are presented as mean ± SD of three independent biological replicates. Asterisks indicate statistically significant differences between treatments as determined by a two-sided Student’s \u003cem\u003et\u003c/em\u003e-test (\u003cem\u003eP \u0026lt; 0.05\u003c/em\u003e).\u003cstrong\u003e g\u003c/strong\u003e-\u003cstrong\u003ei\u003c/strong\u003e Metabolic profiles of the wild‑type (\u003cem\u003ecgdA\u003c/em\u003e::Ctrl), the riboswitch mutant (\u003cem\u003ecgdA\u003c/em\u003e::M2), and the complementary strain (\u003cem\u003ecgdA\u003c/em\u003e::M3) during rehydration. Kinetics of intracellular arginine (\u003cstrong\u003eg\u003c/strong\u003e), guanidine (\u003cstrong\u003eh\u003c/strong\u003e), and succinate (\u003cstrong\u003ei\u003c/strong\u003e) levels were measured during rehydration after three cycles of dehydration-rehydration. Data are shown as mean ± SD from at least four independent biological replicates. Letters above bars denote significant differences (\u003cem\u003eP\u003c/em\u003e \u0026lt; 0.05) assessed by one-way ANOVA across all strains.\u003c/p\u003e","description":"","filename":"Figures3.png","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/1b768c8f756f2034a71a601d.png"},{"id":100869396,"identity":"a7743a57-4592-4332-8f7d-0614058f5457","added_by":"auto","created_at":"2026-01-22 08:59:03","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":297647,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eGuanidine production and degradation pathways exist in \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003eN. flagelliforme\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e.\u003c/strong\u003e \u003cstrong\u003ea\u003c/strong\u003e The schematic illustrates the guanidine production and degradation pathways. Using U-[\u003csup\u003e13\u003c/sup\u003eC,\u003csup\u003e15\u003c/sup\u003eN]-arginine as a tracer, arginine is consumed by the ethylene-forming enzyme (Efe), resulting in \u003csup\u003e15\u003c/sup\u003eN- and \u003csup\u003e13\u003c/sup\u003eC-labeled guanidine and pyrroline-5-carboxylate, while succinate remains unlabeled. The resulting guanidine is subsequently degraded to carboxyguanidine, allophanate, and ultimately to NH\u003csub\u003e3\u003c/sub\u003e and CO\u003csub\u003e2\u003c/sub\u003e, catalyzed sequentially by guanidine carboxylase (GcA), the heterodimeric carboxyguanidine deiminase (CgdA/B), and allophanate hydrolase (AtzF). Atoms enriched with \u003csup\u003e15\u003c/sup\u003eN and \u003csup\u003e13\u003c/sup\u003eC are marked with red and blue symbols, respectively. \u003cstrong\u003eb\u003c/strong\u003e Ion chromatogram of key metabolites from the reaction mixture. \u003cstrong\u003ec\u003c/strong\u003e Mass spectra of selected metabolites, displayed as mass‑to‑charge ratio (m/z). 2-OG, α‑ketoglutarate.\u003c/p\u003e","description":"","filename":"Figures4.png","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/de5ca94cad8c1885108e29a5.png"},{"id":100869395,"identity":"7526f113-3430-4bc5-8e97-22563d614c2e","added_by":"auto","created_at":"2026-01-22 08:59:02","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":463866,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eGuanidine carboxylase pathway is essential for \u003c/strong\u003e\u003cem\u003e\u003cstrong\u003eN. flagelliforme\u003c/strong\u003e\u003c/em\u003e\u003cstrong\u003e recovery from dehydration. a, b \u003c/strong\u003eRepresentative photographs of \u003cem\u003eN. flagelliforme\u003c/em\u003e strains after 6 days of dehydration and rehydration cycles under nitrogen-replete (\u003cstrong\u003ea\u003c/strong\u003e) and nitrogen-deplete (\u003cstrong\u003eb\u003c/strong\u003e) conditions. Cells were pre-adapted for 3 days in the corresponding nitrogen-replete or ‑deplete medium, and 30 mL of culture (OD\u003csub\u003e750\u003c/sub\u003e = 0.5) was collected on nitrocellulose membrane filters for subsequent dehydration-rehydration treatment. Similar results were obtained in three independent experiments.\u003cstrong\u003e c, d\u003c/strong\u003e Growth curves of the wild‑type (\u003cem\u003ecgdA\u003c/em\u003e::Ctrl), riboswitch mutant (\u003cem\u003ecgdA\u003c/em\u003e::M2), and complementary strain (\u003cem\u003ecgdA\u003c/em\u003e::M3) recovered from 6-day dehydration-rehydration cycles under nitrogen-replete (\u003cstrong\u003ec\u003c/strong\u003e) and nitrogen-deplete (\u003cstrong\u003ed\u003c/strong\u003e) conditions. Cultures were recovered in nitrogen-replete medium with an initial OD\u003csub\u003e750\u003c/sub\u003e adjusted to 0.05. Data are presented as mean ± SD of three independent biological replicates. \u003cstrong\u003ee, f\u003c/strong\u003e Representative photographs of different \u003cem\u003eN. flagelliforme\u003c/em\u003e strains recovering from dehydration-rehydration cycles under nitrogen-replete (\u003cstrong\u003ee\u003c/strong\u003e) and nitrogen-deplete (\u003cstrong\u003ef\u003c/strong\u003e) conditions. Experiments were performed in triplicate, with similar results.\u003c/p\u003e","description":"","filename":"Figures5.png","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/5078c5bf5dae734fd6b75531.png"},{"id":100869394,"identity":"bb7e535e-4f7b-43b6-9b94-1402ce65d1ca","added_by":"auto","created_at":"2026-01-22 08:59:02","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":279323,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eThe riboswitch-regulated guanidine carboxylase pathway is widespread among terrestrial cyanobacteria. a Phylogenetic analysis of cyanobacteria harboring the guanidine-I riboswitch.\u003c/strong\u003e The tree was constructed using the maximum-likelihood method based on a concatenated alignment of 31 single-copy protein-coding genes from 60 representative cyanobacterial strains. Branches are color-coded according to taxonomic classification as indicated in the legend. \u003cem\u003eGloeobacter violaceus \u003c/em\u003ePCC 7421 was used as the outgroup. Nodes with bootstrap support values ≥70% are marked with black dots. \u003cstrong\u003eb \u003c/strong\u003eThe diagram shows the arrangement of genes downstream of the guanidine-I riboswitch, with color indicating its frequency within a genome. Solid and open circles denote the presence and absence of the corresponding gene, respectively. Detailed genomic loci of the riboswitch and associated genes are provided in Supplementary Table 3. \u003cstrong\u003ec \u003c/strong\u003eThe frequency with which the guanidine-I riboswitch regulates Gca and GdmH across different environments. Gca\u003csup\u003eP\u003c/sup\u003e and GdmH\u003csup\u003eP\u003c/sup\u003e are defined as the number of genes in the guanidine carboxylase and hydrolase pathways, respectively, under the regulatory control of the guanidine-I riboswitch in a given environment. The combined total of Gca\u003csup\u003eP\u003c/sup\u003e and GdmH\u003csup\u003eP\u003c/sup\u003e genes across the selected environments is 716. Gd-I RS, guanidine-I riboswitch.\u003c/p\u003e","description":"","filename":"Figures6.png","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/5e2ae9d07ff9a67d7a9cdab7.png"},{"id":102298285,"identity":"a18e6fea-9cbe-45d8-a9f4-f06700ec1b81","added_by":"auto","created_at":"2026-02-10 10:36:42","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":2991732,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/44e32d62-ec27-4f71-994a-736f6b38df71.pdf"},{"id":100950448,"identity":"6b085a34-cdcc-4c58-ac51-ee21bb6755ec","added_by":"auto","created_at":"2026-01-23 07:08:13","extension":"xlsx","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":2007646,"visible":true,"origin":"","legend":"Supplementary Data","description":"","filename":"SupplementaryData1.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/79bd847d0c74ca02ac723a13.xlsx"},{"id":100869398,"identity":"d99672f0-73a9-44b6-aaec-541e63e052b7","added_by":"auto","created_at":"2026-01-22 08:59:03","extension":"pdf","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":5824981,"visible":true,"origin":"","legend":"Supplementary Figures 1-15","description":"","filename":"SupplementaryFigures.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/a562e99704b8515625a1c601.pdf"},{"id":100869397,"identity":"4b18173d-bf38-4f73-a614-52bcf9d4150f","added_by":"auto","created_at":"2026-01-22 08:59:03","extension":"xlsx","order_by":3,"title":"","display":"","copyAsset":false,"role":"supplement","size":275524,"visible":true,"origin":"","legend":"Supplementary Tables S1\u0026#x2013;S5","description":"","filename":"SupplementaryTableS1XXXS5.xlsx","url":"https://assets-eu.researchsquare.com/files/rs-8507520/v1/20bf076f6c42c571af6177cf.xlsx"}],"financialInterests":"There is \u003cb\u003eNO\u003c/b\u003e Competing Interest.","formattedTitle":"Guanidine fuels rapid resurrection of desert cyanobacteria","fulltext":[{"header":"Introduction","content":"\u003cp\u003eDesert ecosystems are characterized by extreme aridity, where microbial activity is largely restricted to brief and unpredictable hydration pulses\u003csup\u003e1\u0026ndash;3\u003c/sup\u003e. The rapid resuscitation of soil microbial communities following rewetting is crucial for ecosystem functioning, yet this process is consistently limited by nitrogen (N) availability\u003csup\u003e3,4\u003c/sup\u003e. Although biological nitrogen fixation represents the primary nitrogen source in these environments\u003csup\u003e5\u003c/sup\u003e, the regulatory mechanisms governing nitrogen cycling and microbial resuscitation dynamics remain poorly understood.\u003c/p\u003e\n\u003cp\u003eCyanobacteria, as pioneer organisms in these habitats, face a critical metabolic challenge\u003csup\u003e6,7\u003c/sup\u003e. They must rapidly synthesize nitrogen-intensive cellular components during brief periods of moisture and light availability\u003csup\u003e2,8\u003c/sup\u003e. However, biological nitrogen fixation is inefficient in this initial phase due to its high energy demands and the delayed activation of the nitrogenase complex\u003csup\u003e9,10\u003c/sup\u003e. This contradiction between an immediate high nitrogen demand and a limited supply suggests that the efficient remobilization of internal nitrogen reserves is essential for survival.\u003c/p\u003e\n\u003cp\u003eSeveral molecules are involved in nitrogen storage and metabolism in cyanobacteria, with cyanophycin serving as a major intracellular nitrogen reservoir\u003csup\u003e11,12\u003c/sup\u003e. This non-ribosomally synthesized copolymer is broken down into aspartate and arginine under nitrogen starvation\u003csup\u003e11,12\u003c/sup\u003e. ArgZ (AgrE), a bifunctional enzyme, then metabolizes arginine via its arginine dihydrolase activity, converting it to ornithine\u0026nbsp;while releasing CO\u003csub\u003e2\u003c/sub\u003e and ammonia\u003csup\u003e13,14\u003c/sup\u003e. The resulting ornithine can either enter nitrogen assimilation via the ornithine\u0026ndash;ammonia cycle or be converted to proline by the ornithine cyclodeaminase activity of AgrE, followed by eventual degradation via the proline oxidase PutA\u003csup\u003e13,15\u003c/sup\u003e. Through the combined AgrE/PutA pathway, the four nitrogen atoms of arginine are converted into three molecules of ammonia and one molecule of glutamate, thereby efficiently remobilizing nitrogen for metabolic use\u003csup\u003e15\u003c/sup\u003e.\u003c/p\u003e\n\u003cp\u003eRecent studies have\u0026nbsp;significantly advanced our understanding of nitrogen metabolism, demonstrating that some microorganisms can utilize guanidine and its derivatives\u0026nbsp;as nitrogen sources\u003csup\u003e16\u0026ndash;23\u003c/sup\u003e. Guanidine, a chemically\u0026nbsp;stable\u0026nbsp;compound containing 71.1% nitrogen by mass, was discovered more than 150 years\u0026nbsp;ago\u003csup\u003e24\u003c/sup\u003e. However, the pathways and mechanisms underlying its biodegradation remain poorly understood. It is now known that microbes degrade guanidine primarily via two routes: the\u0026nbsp;guanidinase\u0026nbsp;pathway and the\u0026nbsp;carboxylase pathway\u003csup\u003e17,20,21\u003c/sup\u003e. Both pathways ultimately convert one molecule of guanidine into three molecules of ammonia. While certain cyanobacteria and complete ammonia oxidizers (comammox) have been shown to\u0026nbsp;utilize\u0026nbsp;guanidine as the sole nitrogen source through\u0026nbsp;the guanidinase pathway\u003csup\u003e20,21\u003c/sup\u003e, the physiological role of the guanidine carboxylase pathway remains unclear in cyanobacteria, and its ecological significance warrants further investigation.\u003c/p\u003e\n\u003cp\u003eHere, we examined nitrogen remobilization mediated by the guanidine carboxylase pathway in the\u0026nbsp;representative\u0026nbsp;desert cyanobacterium\u0026nbsp;\u003cem\u003eNostoc flagelliforme\u003c/em\u003e (hereafter \u003cem\u003eN. flagelliforme\u003c/em\u003e). Our results revealed that this pathway is regulated by a guanidine-I riboswitch in \u003cem\u003eN. flagelliforme\u003c/em\u003e, a mechanism likely conserved across many other terrestrial cyanobacteria. The presence of this riboswitch-controlled system may enable desert cyanobacteria to rapidly mobilize internal nitrogen reserves during rehydration, thereby facilitating their post-desiccation recovery. This capability has important implications for biogeochemical cycling and ecosystem stability in arid and semi-arid regions.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003e\u003cstrong\u003eA guanidine-binding riboswitch is located within the 5\u0026prime; UTR of the \u003cem\u003ecgdA-cgdB-gcA-atzF\u0026nbsp;\u003c/em\u003ecluster in\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eN. flagelliforme\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e.\u0026nbsp;\u003c/strong\u003e\u003cem\u003eN. flagelliforme\u003c/em\u003e is\u0026nbsp;a model desert cyanobacterium renowned for its extreme desiccation tolerance and rapid recovery upon rehydration\u003csup\u003e10,25\u003c/sup\u003e.\u0026nbsp;The\u0026nbsp;\u003cem\u003ecgdA-cgdB-gcA-atzF\u003c/em\u003e gene cluster, implicated in urea degradation (Fig. 1a), was previously identified from transcriptomic data as being\u0026nbsp;differentially expressed during rehydration\u003csup\u003e10\u003c/sup\u003e.\u0026nbsp;RT‑qPCR analysis confirmed that the transcription of\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e,\u0026nbsp;\u003cem\u003ecgdB\u003c/em\u003e,\u0026nbsp;\u003cem\u003egcA\u003c/em\u003e, and\u0026nbsp;\u003cem\u003eatzF\u003c/em\u003e was significantly upregulated in the early phase of rehydration (Supplementary Fig. 1a). A similar induction pattern was also observed in two dominant biological soil crust cyanobacteria,\u0026nbsp;\u003cem\u003eMicrocoleus vaginatus\u003c/em\u003e (\u003cem\u003eM. vaginatus\u003c/em\u003e,\u0026nbsp;Supplementary Fig. 1b)\u003csup\u003e26\u003c/sup\u003e and\u0026nbsp;\u003cem\u003eLeptolyngbya ohadii\u003c/em\u003e (\u003cem\u003eL. ohadii\u003c/em\u003e,\u0026nbsp;Supplementary Fig. 1c)\u003csup\u003e1\u003c/sup\u003e, suggesting a conserved role of this gene cluster during rehydration across desiccation-tolerant cyanobacteria.\u003c/p\u003e\n\u003cp\u003eBioinformatic analysis\u0026nbsp;identified\u0026nbsp;a putative guanidine riboswitch within the 5\u0026prime; untranslated region (UTR) of this gene cluster (Fig. 1b). Secondary structure modeling\u0026nbsp;revealed\u0026nbsp;that\u0026nbsp;it adopts the characteristic guanidine-I architecture, featuring\u0026nbsp;two stem loops (P1 and P2) and a long 3\u0026prime; tail overlapping a terminator hairpin (P3, Fig. 1b),\u0026nbsp;consistent with known riboswitches\u0026nbsp;in\u0026nbsp;\u003cem\u003eSulfobacillus acidophilus\u003c/em\u003e\u003csup\u003e27\u003c/sup\u003e and\u0026nbsp;\u003cem\u003eDickeya dadantii\u003c/em\u003e\u003csup\u003e28\u003c/sup\u003e.\u003c/p\u003e\n\u003cp\u003eTo\u0026nbsp;assess\u0026nbsp;the substrate-binding ability of this RNA aptamer, isothermal titration calorimetry (ITC) was performed using guanidine hydrochloride as the titrant (Fig. 1d-g). The\u0026nbsp;data revealed that the 5\u0026prime; UTR of \u003cem\u003eN. flagelliforme\u003c/em\u003e \u003cem\u003ecgdA\u003c/em\u003e binds guanidine with high affinity (Fig. 1d), but not urea\u0026nbsp;(Supplementary Fig. 2). The dissociation constant (\u003cem\u003eK\u003c/em\u003e\u003csub\u003ed\u003c/sub\u003e) for guanidine was determined to be\u0026nbsp;26 \u0026micro;M (Fig. 1d), which is lower than\u0026nbsp;the previously reported value of\u0026nbsp;39.2 \u0026plusmn; 0.85 \u0026mu;M in\u0026nbsp;\u003cem\u003eD. dadantii\u003c/em\u003e\u003csup\u003e28\u003c/sup\u003e. To\u0026nbsp;exclude nonspecific\u0026nbsp;binding of the RNA aptamer to guanidine,\u0026nbsp;we introduced mutations into the P1 domain of the riboswitch\u0026nbsp;(Fig. 1c).\u0026nbsp;Deletion of the predicted\u0026nbsp;guanidine-Ⅰ riboswitch (M1) abolished guanidine binding in the ITC assays (Fig. 1e), as did\u0026nbsp;the disruption of the conserved\u0026nbsp;base-pairing in mutant M2 (Fig. 1f).\u0026nbsp;In contrast, binding was restored with\u0026nbsp;the compensatory mutation M3 (Fig. 1g).\u0026nbsp;Together, these results\u0026nbsp;demonstrate that the 5\u0026prime; UTR of the\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e gene contains\u0026nbsp;a functional riboswitch that specifically recognizes guanidine.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eThe guanidine riboswitch regulates expression of the \u003cem\u003ecgdA-cgdB-gcA-atzF\u0026nbsp;\u003c/em\u003egene cluster.\u0026nbsp;\u003c/strong\u003eBioinformatic prediction of an NtcA binding site (5\u0026prime;-GTAgtaagtgaTAC-3\u0026prime;) within the \u003cem\u003ecgdA\u003c/em\u003e promoter region (Supplementary Fig. 3) indicated that the expression of \u003cem\u003ecgdA\u003c/em\u003e in \u003cem\u003eN. flagelliforme\u003c/em\u003e is likely regulated by the global nitrogen regulator NtcA\u003csup\u003e29\u0026ndash;31\u003c/sup\u003e. This interaction was validated by an electrophoretic mobility shift assay (EMSA; Supplementary Fig. 3c). To investigate the function of the guanidine riboswitch \u003cem\u003ein vivo\u003c/em\u003e, site-directed mutations were introduced into its native locus using CRISPR-Cpf1 genome editing\u003csup\u003e32\u003c/sup\u003e (Fig. 2a-c). RT-qPCR analysis revealed that in wild-type background controls (\u003cem\u003ecgdA::\u003c/em\u003eCtrl), genes downstream of the promoter were significantly induced under nitrate starvation (Fig. 2a), consistent with the typical expression pattern of the NtcA regulon.\u0026nbsp;Notably, this induction was abolished in the \u003cem\u003ecgdA::\u003c/em\u003eM2 mutant (Fig. 2b). The loss of induction was specifically attributable to the disruption of the riboswitch and not to any other effects, as evidenced by the restored gene induction in the compensatory mutant \u003cem\u003ecgdA\u003c/em\u003e::M3 (Fig. 2c).\u003c/p\u003e\n\u003cp\u003eThe transcriptional response of the\u0026nbsp;\u003cem\u003ecgdA-cgdB-gcA-atzF\u003c/em\u003e gene cluster\u0026nbsp;was further assessed\u0026nbsp;using a luciferase reporter system. In this\u0026nbsp;assay,\u0026nbsp;promoter activity was\u0026nbsp;measured by quantifying\u0026nbsp;LuxAB-derived bioluminescence\u0026nbsp;following fusion to different promoter variants, each of which retained the NtcA-binding site.\u0026nbsp;Compared to cultures grown under nitrogen-replete conditions (BG11, day 0), reporter strains carrying the wild-type\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e promoter\u0026nbsp;exhibited a rapid increase\u0026nbsp;in LuxAB bioluminescence during the first three days of incubation in\u0026nbsp;nitrate-depleted medium\u0026nbsp;(BG11\u003csub\u003e0\u003c/sub\u003e; Fig. 2d).\u0026nbsp;Although\u0026nbsp;the NtcA-binding site\u0026nbsp;remained intact,\u0026nbsp;no increase in\u0026nbsp;bioluminescence\u0026nbsp;was observed in strains with\u0026nbsp;an inactivated guanidine riboswitch\u0026nbsp;(M2; Fig. 2e).\u0026nbsp;In contrast, introducing\u0026nbsp;a compensatory mutation (M3) restored nitrogen-starvation-induced\u0026nbsp;expression of\u003cem\u003e\u0026nbsp;cgdA\u003c/em\u003e (Fig. 2f).\u0026nbsp;Together, these findings underscore the physiological relevance of the\u0026nbsp;\u003cem\u003ecgdA-cgdB-gcA-atzF\u003c/em\u003e gene cluster in the low-N response, as well as the essential role of the guanidine riboswitch in regulating it.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eThe expression of the \u003cem\u003ecgdA-cgdB-gcA-atzF\u0026nbsp;\u003c/em\u003ecluster is also regulated by dehydration and rehydration.\u0026nbsp;\u003c/strong\u003eThe transcriptional dynamics of the \u003cem\u003ecgdA\u003c/em\u003e-\u003cem\u003ecgdB\u003c/em\u003e-\u003cem\u003egcA\u003c/em\u003e-\u003cem\u003eatzF\u003c/em\u003e gene cluster during dehydration and rehydration cycles were further examined using the same luciferase reporter strains (Fig. 3a-f). In this assay, \u003cem\u003eN. flagelliforme\u003c/em\u003e cells were first cultured for 3 days in liquid BG11 or BG11\u003csub\u003e0\u003c/sub\u003e medium, collected on a nitrocellulose membrane, and subjected to periodic dehydration and rehydration cycles (21 h: 3 h). Dehydration was achieved by air-drying membranes at 70% relative humidity, while rehydration was performed by placing membranes on 1.5% (m/v) BG11 agar plates with or without nitrate\u003csup\u003e33\u003c/sup\u003e. As anticipated, nitrogen-starved reporter strains carrying either the wild-type (Fig. 3a) or the compensatory M3 (Fig. 3c) riboswitch exhibited stronger luminescence during the dehydration and rehydration cycles. In contrast, the luminescence signal from the nitrogen-depleted M2-\u003cem\u003eluxAB\u003c/em\u003e strain remained similar to that observed under nitrogen-replete conditions (Fig. 3b).\u003c/p\u003e\n\u003cp\u003eTo comprehensively assess\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e promoter activity under fluctuating water availability, time-course bioluminescence analysis was performed on day 6 (the third dehydration-rehydration cycle). As shown in Fig. 3d, bioluminescence increased during rehydration and decreased during dehydration. Notably, this oscillatory expression pattern was not of circadian origin. When rehydration was extended by an additional three hours, \u003cem\u003ecgdA\u003c/em\u003e promoter activity remained elevated and only decreased upon re-exposure to dehydration (Supplementary Fig. 4). Consistently, bioluminescence induction was absent in the guanidine riboswitch-inactivated strain (M2; Fig. 3e) but was restored in the compensatory mutant (M3; Fig. 3f). These results indicate that the guanidine riboswitch is crucial for activating the \u003cem\u003ecgdA\u003c/em\u003e-\u003cem\u003ecgdB\u003c/em\u003e-\u003cem\u003egcA\u003c/em\u003e-\u003cem\u003eatzF\u003c/em\u003e gene cluster during rehydration of nitrogen-starved \u003cem\u003eN. flagelliforme\u003c/em\u003e.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eThe \u003cem\u003ecgdA-cgdB-gcA-atzF\u0026nbsp;\u003c/em\u003ecluster is responsible for guanidine carboxylation in\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eN. flagelliforme\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e.\u0026nbsp;\u003c/strong\u003eWhile genome annotations have linked the\u0026nbsp;\u003cem\u003ecgdA-cgdB-gcA-atzF\u0026nbsp;\u003c/em\u003ecluster to the urea carboxylation process, a recent study in gamma proteobacteria suggested\u0026nbsp;its potential involvement in guanidine carboxylation\u003csup\u003e17\u003c/sup\u003e.\u0026nbsp;Consistent with this,\u0026nbsp;our ITC\u0026nbsp;data demonstrated that\u0026nbsp;the 5\u0026prime; UTR of\u0026nbsp;this cluster\u0026nbsp;specifically binds guanidine but not urea,\u0026nbsp;indicating its role in guanidine metabolism.\u0026nbsp;To\u0026nbsp;test whether a functional\u0026nbsp;guanidine carboxylation pathway exists in\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e,\u0026nbsp;we employed\u0026nbsp;U-[\u003csup\u003e13\u003c/sup\u003eC,\u003csup\u003e15\u003c/sup\u003eN]-arginine as a tracer. In the presence of ethylene-forming enzyme (Efe) performing a reaction as described for the\u0026nbsp;\u003cem\u003ePseudomonas savastanoi\u003c/em\u003e Efe (Supplementary Fig. 5)\u003csup\u003e34\u003c/sup\u003e, arginine should be converted to guanidine labeled accordingly (Fig. 4). LC-MS\u0026nbsp;analysis successfully\u0026nbsp;detected \u003csup\u003e15\u003c/sup\u003eN- and \u003csup\u003e13\u003c/sup\u003eC-labeled guanidine, as well as its\u0026nbsp;isotopically\u0026nbsp;labeled degradation products, carboxyguanidine and allophanate (Fig. 4),\u0026nbsp;providing direct evidence for the presence of an\u0026nbsp;arginine-to-guanidine Efe activity and of\u0026nbsp;an active\u0026nbsp;guanidine carboxylation pathway in\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e.\u003c/p\u003e\n\u003cp\u003eWe also compared the metabolism of the wild-type\u0026nbsp;(\u003cem\u003ecgdA\u003c/em\u003e::Ctrl), riboswitch mutant\u0026nbsp;(\u003cem\u003ecgdA\u003c/em\u003e::M2),\u0026nbsp;and complementary strain\u0026nbsp;(\u003cem\u003ecgdA\u003c/em\u003e::M3)\u0026nbsp;during the rewetting phase following cycles of dehydration and rehydration. After three days of periodic\u0026nbsp;dehydration and rehydration, the\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::M2 mutant\u0026nbsp;accumulated significantly less intracellular arginine (Fig. 3g). Interestingly, arginine levels in this mutant increased upon rehydration (Fig. 3g), likely due to nitrogen remobilization via the cyanophycin breakdown pathway. In contrast, such arginine accumulation was not observed in\u0026nbsp;the\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::Ctrl and\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::M3 strains (Fig. 3g), suggesting that in these strains, arginine was rapidly converted into guanidine and succinate by the guanidine biosynthesis protein Efe. This was supported by measurements of guanidine and succinate levels\u0026nbsp;(Fig. 3 h-i). Because the\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::M2\u0026nbsp;strain could not induce guanidine carboxylation genes during rehydration, it accumulated substantially more guanidine than the\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::Ctrl and\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::M3 strains (Fig. 3h). Together, these results indicate that\u0026nbsp;the\u0026nbsp;\u003cem\u003ecgdA-cgdB-gcA-atzF\u003c/em\u003e cluster\u0026nbsp;is essential for guanidine breakdown by \u003cem\u003eN. flagelliforme\u003c/em\u003e.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eGuanidine carboxylation\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eplays an essential role in adaptation to dehydration and rehydration\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;cycles.\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003eTo investigate the physiological role of the guanidine carboxylation pathway, we measured the photosynthetic performance of \u003cem\u003eN. flagelliforme\u003c/em\u003e strains (\u003cem\u003ecgdA\u003c/em\u003e::Ctrl, \u003cem\u003ecgdA\u003c/em\u003e::M2, and \u003cem\u003ecgdA\u003c/em\u003e::M3) after 3 hours of recovery following successive dehydration and rehydration cycles. After three days of treatment, the \u003cem\u003ecgdA::\u003c/em\u003eM2 mutant exhibited significantly lower rates of oxygen evolution and respiration (Supplementary Fig. 6a). The phycobilisome absorbance in the \u003cem\u003ecgdA::\u003c/em\u003eM2 strain was also lower than that in the \u003cem\u003ecgdA\u003c/em\u003e::Ctrl and \u003cem\u003ecgdA\u003c/em\u003e::M3 strains (Supplementary Fig. 6b). Additionally, nitrogenase activity in \u003cem\u003ecgdA\u003c/em\u003e::M2 was lower than that in the other two strains (Supplementary Fig. 6c). These diminished metabolic activities likely resulted from the impaired recovery of the \u003cem\u003ecgdA::\u003c/em\u003eM2 strain during rehydration, as time-series chlorophyll fluorescence measurements showed that this riboswitch-inactivated mutant exhibited slower photosynthetic recovery, especially after multiple rounds of dehydration and rehydration (Supplementary Fig. 6d).\u003c/p\u003e\n\u003cp\u003eTo further understand the transcriptional responses during recovery, we analyzed the transcriptomic profiles of\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e following successive dehydration\u0026ndash;rehydration cycles. Based on Fuzzy C-Means clustering (FCM;\u0026nbsp;Supplementary Fig. 7), genes associated with ribosomal function, DNA replication, recombination, and repair were upregulated in the\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e::M2 strain during rehydration (Cluster 2; Supplementary Table 1). In contrast, genes involved in photosynthesis, nitrogen fixation, hydrogenase activity, and the guanidine carboxylation pathway were downregulated (Cluster 4; Supplementary Table 1 and Supplementary Figs. 7-9), consistent with the physiological differences observed between \u003cem\u003ecgdA\u003c/em\u003e::Ctrl and \u003cem\u003ecgdA\u003c/em\u003e::M2 strains (Supplementary Fig. 6). Further analysis showed that the nitrogen‑limitation biomarker genes \u003cem\u003enrtA\u003c/em\u003e (FC=1.56, \u003cem\u003eP\u003c/em\u003e\u0026lt;0.05), \u003cem\u003enrtB\u003c/em\u003e (FC=2.33, \u003cem\u003eP\u003c/em\u003e\u0026lt;0.05), and \u003cem\u003eamt1\u003c/em\u003e (FC=1.73, \u003cem\u003eP\u003c/em\u003e\u0026lt;0.05) were more highly expressed in \u003cem\u003ecgdA\u003c/em\u003e::M2 (Supplementary Data 1), indicating a more constrained nitrogen supply in the riboswitch‑inactivated mutant. Consistent with this, the quantification of intracellular cyanophycin revealed that \u003cem\u003eN. flagelliforme\u003c/em\u003e\u003cem\u003e\u0026nbsp;\u003c/em\u003estrains unable to induce guanidine carboxylase (M2) exhibited reduced cyanophycin degradation efficiency (Supplementary Fig. 10). Together, these results demonstrate that disruption of the guanidine carboxylation pathway significantly impairs nitrogen remobilization during rehydration.\u003c/p\u003e\n\u003cp\u003eAfter six days of periodic dehydration and rehydration, \u003cem\u003ecgdA\u003c/em\u003e::M2 strains grown under nitrogen-replete conditions exhibited a growth pattern similar to that of \u003cem\u003ecgdA\u003c/em\u003e::Ctrl and \u003cem\u003ecgdA\u003c/em\u003e::M3 strains (Fig. 5a). In contrast, under nitrogen-deprived conditions, the \u003cem\u003ecgdA\u003c/em\u003e::M2 strain accumulated less biomass and displayed more pronounced chlorosis than the other two strains following the same treatment (Fig. 5b), despite the higher expression of \u003cem\u003eargZ\u003c/em\u003e in the mutant (Supplementary Data 1). When recovered in liquid BG11 medium (Fig. 5c-f), only the nitrogen-starved \u003cem\u003ecgdA\u003c/em\u003e::Ctrl and \u003cem\u003ecgdA\u003c/em\u003e::M3 strains were able to rapidly resume growth and regain green pigmentation after six days, whereas the resuscitation rate of nitrogen-starved \u003cem\u003ecgdA\u003c/em\u003e::M2 was markedly lower (Fig. 5d, f). These results indicate that the riboswitch-regulated guanidine carboxylation pathway is essential for cell survival and recovery during dehydration and rehydration cycles.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eThe guanidine carboxylation gene cluster originated from an ancient horizontal gene transfer event.\u0026nbsp;\u003c/strong\u003eTo elucidate the evolutionary origin of the cyanobacterial guanidine carboxylation pathway, we\u0026nbsp;assembled a comprehensive dataset comprising 16S rDNA sequences and concatenated CgdA and CgdB protein sequences. Phylogenetic reconstruction revealed that the maximum-likelihood species tree was largely congruent with the guanidine catabolism tree inferred from the concatenated CgdA and CgdB alignment (Supplementary Figs. 11, 12). All cyanobacterial guanidine carboxylation genes formed a distinct monophyletic clade separate from those of other bacteria (Supplementary Fig. 12), implying that cyanobacteria likely acquired an ancestral carboxylation module for guanidine degradation from proteobacteria via horizontal gene transfer (HGT)\u003csup\u003e10,35\u003c/sup\u003e, followed by gene duplication and functional diversification into different cyanobacterial lineages. Moreover, phylogenetic comparisons indicate that the marker genes have coevolved with the guanidine‑degradation genes in cyanobacteria, as supported by their high topological congruence (Supplementary Fig. 13). Together, these results suggest that the HGT‑mediated acquisition of an ancestral carboxylation reaction, which initiates guanidine degradation, likely facilitated the adaptive radiation of these cyanobacteria into dynamic habitats.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eThe riboswitch-regulated guanidine carboxylase pathway is predominantly found in terrestrial cyanobacteria.\u0026nbsp;\u003c/strong\u003eTo\u0026nbsp;examine the distribution of\u0026nbsp;the riboswitch-regulated guanidine carboxylase pathway in cyanobacteria, we\u0026nbsp;performed\u0026nbsp;comparative genomic analyses of strains from\u0026nbsp;diverse representative habitats.\u0026nbsp;Homology searches across 2020 cyanobacterial strains identified 835 unique guanidine-I RNA sequence motifs (RF00442 in the Rfam database) distributed among\u0026nbsp;739 strains (Supplementary Table 2 and Supplementary Fig. 14). While most of these cyanobacteria contain a single copy of the guanidine-I riboswitch, some strains carry two or three distinct copies (Fig. 6 and Supplementary Table 2). These results indicate that the guanidine-I riboswitches are widely distributed across the cyanobacterial phylum.\u003c/p\u003e\n\u003cp\u003eTo identify genes linked to the guanidine‑I aptamer, we compared the sequences immediately downstream of the aptamer with the NCBI protein database. Among the 739 genomes analyzed, the aptamer was located upstream of\u0026nbsp;\u003cem\u003egdmH\u003c/em\u003e (encoding guanidine hydrolase\u003csup\u003e20\u003c/sup\u003e) in 648 cases and upstream of\u0026nbsp;\u003cem\u003ecgdAB\u003c/em\u003e (encoding carboxyguanidine deiminase) in 91 cases (Supplementary Table 2). Intriguingly, although both pathways are involved in guanidine utilization, they appear to be mutually exclusive, as no co-occurrence of the two has been observed. Further analysis revealed that the guanidine‑I riboswitch-regulated carboxylase pathway was predominantly distributed in terrestrial cyanobacteria (Fig. 6c and Supplementary Table 2). Moreover, its presence shows a positive association with habitat aridity (Fig. 6c).\u003c/p\u003e"},{"header":"Discussion","content":"\u003cp\u003eDesert ecosystems exhibit low nitrogen bioavailability, due to limited external inputs and incomplete nitrogen cycling\u003csup\u003e36\u0026ndash;38\u003c/sup\u003e. This nitrogen limitation constrains microbial activity, shapes the soil community structure, and ultimately restricts primary productivity. To survive such extremes, desert cyanobacteria must rapidly remobilize intracellular nitrogen when water becomes available. However, the mechanism by which they acquire usable nitrogen during rehydration has remained enigmatic. Here, we present multiple evidence that the riboswitch-controlled guanidine carboxylase pathway is essential for nitrogen remobilization upon rehydration. Although the\u0026nbsp;hydrolysis pathway appears to be more common in aquatic cyanobacteria, the carboxylase pathway is ecologically significant in terrestrial habitats. This efficient nitrogen-remobilization pathway likely enables desert cyanobacteria to swiftly restore cellular functions during the brief periods when both water and light are available, thus priming them for subsequent dehydration.\u003c/p\u003e\n\u003cp\u003eSeveral metabolic pathways have been implicated in the mobilization of intracellular nitrogen reserves in cyanobacteria. Transcriptomic analysis revealed high expression of\u0026nbsp;\u003cem\u003eagrE\u003c/em\u003e (\u003cem\u003eargZ\u003c/em\u003e) and \u003cem\u003eputA\u003c/em\u003e in both \u003cem\u003ecgdA::\u003c/em\u003eCtrl and \u003cem\u003ecgdA::\u003c/em\u003eM2 strains (Supplementary Data 1), underscoring their importance in nitrogen remobilization during the rehydration process. Although the AgrE/PutA pathway is highly efficient\u003csup\u003e15\u003c/sup\u003e, our data show that \u003cem\u003eN. flagelliforme\u003c/em\u003e strains unable to upregulate the guanidine carboxylase pathway (\u003cem\u003ecgdA\u003c/em\u003e::M2) displayed significantly reduced cellular activity and slower photosynthetic recovery, even though \u003cem\u003eagrE\u003c/em\u003e expression was higher than in the wild‑type (Supplementary Fig. 9). These results indicate that arginine catabolism via the AgrE/PutA pathway alone cannot meet the high nitrogen demand of rehydrating cyanobacteria, and further highlight the guanidine carboxylase pathway as a critical mechanism for supporting elevated nitrogen requirements and fueling cellular recovery.\u003c/p\u003e\n\u003cp\u003eMoreover, in contrast to arginine remobilization via the AgrE/PutA pathway and the arginine‑ammonia cycle\u003csup\u003e13,15\u003c/sup\u003e, the Efe-catalyzed conversion of arginine to guanidine consumes three molecules of \u0026alpha;-ketoglutarate (2-OG; Fig. 4a)\u003csup\u003e34\u003c/sup\u003e. Given the central role of 2‑OG in regulating carbon-nitrogen metabolic balance and its marked accumulation under nitrogen-limited conditions\u003csup\u003e39\u0026ndash;42\u003c/sup\u003e, guanidine-dependent arginine degradation may provide a route for nitrogen-starved cyanobacteria to simultaneously catabolize stored carbon and rapidly restore metabolic homeostasis.\u003c/p\u003e\n\u003cp\u003eWhile the role of NtcA in nitrogen homeostasis is widely recognized\u003csup\u003e29\u0026ndash;31\u003c/sup\u003e, our findings elucidate an additional layer of regulation mediated by the guanidine‑I riboswitch for\u0026nbsp;guanidine carboxylation and degradation. ITC analysis confirmed that the guanidine‑I riboswitch in the \u003cem\u003eN. flagelliforme\u003c/em\u003e carboxylase gene cluster specifically binds guanidine, but not urea, validating the precision of this metabolite‑dependent switch. Notably, homologous riboswitches are commonly found upstream of guanidine carboxylase or hydrolase clusters in diverse cyanobacterial strains (Supplementary Table 2). This consistent genomic association suggests that the riboswitch‑dependent regulatory mechanism described here is likely conserved across cyanobacteria.\u003c/p\u003e\n\u003cp\u003eInterestingly, analysis of guanidine carboxylase gene distribution shows that guanidine‑I riboswitch‑controlled carboxylase pathways are primarily found in terrestrial cyanobacteria\u0026nbsp;(Fig. 6c). This pattern matches the ecological niche of\u0026nbsp;\u003cem\u003ePseudomonas syringae\u003c/em\u003e\u003csup\u003e17\u003c/sup\u003e, the model organism for guanidine carboxylation, but differs markedly from that of nickel (Ni)-dependent guanidine hydrolase genes, which are more frequently found in aquatic environments (Fig. 6c). Although Ni is generally scarce in both terrestrial and aquatic environments, its concentration in the ocean exceeds that of other essential trace metals, such as Cu, Fe, and Mn\u003csup\u003e43,44\u003c/sup\u003e. In marine systems, low Ni availability does not appear to drive the loss of Ni-dependent pathways. Instead, marine cyanobacteria preferentially utilize Ni-dependent superoxide dismutase (NiSOD) over SODs that rely on other metal cofactors\u003csup\u003e45,46\u003c/sup\u003e. In contrast, many terrestrial ecosystems exhibit particularly low Ni bioavailability\u003csup\u003e47,48\u003c/sup\u003e. In alkaline soils of arid and semi-arid regions, where precipitation-driven acidification and leaching are minimal, Ni deficiency is likely widespread. Under these conditions, replacing Ni-dependent guanidine hydrolase with a carboxylase pathway may have relieved desert cyanobacteria of the metabolic burden of acquiring scarce nickel\u003csup\u003e17,20,21\u003c/sup\u003e.\u003c/p\u003e\n\u003cp\u003eMoreover, the Michaelis constant (\u003cem\u003eK\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e) of\u0026nbsp;\u003cem\u003ePseudomonas syringae\u003c/em\u003e GcA for guanidine is 0.21 \u0026plusmn; 0.09 mM\u003csup\u003e17\u003c/sup\u003e, which is approximately an order of magnitude lower than the values reported for guanidinase in \u003cem\u003eSynechocystis\u003c/em\u003e sp. PCC 6803 (7.81 \u0026plusmn; 1.80 mM)\u003csup\u003e20\u003c/sup\u003e and \u003cem\u003eNitrospira inopinata\u003c/em\u003e (13.6\u0026thinsp; \u0026plusmn; 0.76 mM)\u003csup\u003e21\u003c/sup\u003e. This lower \u003cem\u003eK\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e of the carboxylase at physiological pH reflects a high catalytic efficiency, enabling effective guanidine utilization even at low substrate concentrations during rehydration.\u003c/p\u003e\n\u003cp\u003eRecent studies have significantly advanced our understanding of guanidine biosynthesis\u003csup\u003e49\u003c/sup\u003e. In addition to the reaction catalyzed by Efe, microorganisms are known to produce guanidine\u0026nbsp;during naphthyridinomycin biosynthesis and through the complete degradation of metformin, one of the world\u0026rsquo;s most widely prescribed drugs\u003csup\u003e23,50\u0026ndash;52\u003c/sup\u003e. In land plants, homoarginine‑6‑hydroxylases are estimated to generate approximately\u0026nbsp;3.5 \u0026times; 10\u003csup\u003e5\u003c/sup\u003e tons of guanidine annually\u003csup\u003e53\u003c/sup\u003e. However, plants appear to primarily transport rather than metabolize guanidine\u003csup\u003e54\u003c/sup\u003e, suggesting that its accumulation or secretion may serve as a strategy to modulate associated microbial communities\u003csup\u003e55\u003c/sup\u003e. Intriguingly, our analysis indicated that some terrestrial cyanobacteria lacking known guanidine‑producing genes harbor gene clusters encoding ATP‑binding cassette (ABC)‑type guanidine transporters (Fig. 6). For instance, \u003cem\u003eMicrocoleus vaginatus\u003c/em\u003e lacks identified guanidine synthesis genes, yet it encodes both the guanidine carboxylation pathway and a high‑affinity guanidine importer, and harbors a guanidine‑I riboswitch upstream of its ABC transporter locus (Supplementary Fig. 15). \u003cem\u003eMicrocoleus\u003c/em\u003e\u003cem\u003e\u0026nbsp;\u003c/em\u003especies are representative biocrust cyanobacteria that do not fix dinitrogen yet dominate nitrogen‑limited biological soil crusts\u003csup\u003e26\u003c/sup\u003e. Although nutrient exchange with associated microbes is well documented in \u003cem\u003eMicrocoleus\u003c/em\u003e and urea has been proposed as the main vehicle for interspecies nitrogen transfer\u003csup\u003e56\u0026ndash;59\u003c/sup\u003e, our analysis suggests that its annotated urea carboxylase is more likely to function in guanidine carboxylation (Supplementary Fig. 15c). Thus, guanidine-based mutualistic nitrogen transfer within biocrusts merits further investigation.\u003c/p\u003e\n\u003cp\u003eCollectively, our findings shed light on the physiological role of guanidine catabolism during rehydration. The guanidine‑I riboswitch controlled carboxylation pathway exhibits high nitrogen remobilization efficiency and restricted phylogenetic distribution outside bacteria\u003csup\u003e20\u003c/sup\u003e. This indicates that it gives \u003cem\u003eN. flagelliforme\u003c/em\u003e\u003cem\u003e\u0026nbsp;\u003c/em\u003ea competitive advantage in nitrogen-poor deserts. This metabolic adaptation could be critical for cyanobacterial survival in these environments, with broader implications for primary productivity and biogeochemical cycling in dryland ecosystems.\u003c/p\u003e"},{"header":"Materials and methods","content":"\u003cp\u003e\u003cstrong\u003eCyanobacterial strains and culture conditions.\u003c/strong\u003e The axenic strain\u0026nbsp;\u003cem\u003eNostoc flagelliforme\u003c/em\u003e CCNUN1 was isolated from\u0026nbsp;dried field samples collected in Sunitezuoqi, Inner Mongolia, China\u003csup\u003e60\u003c/sup\u003e. All strains used in this study are listed in Supplementary Table 5.\u0026nbsp;Unless otherwise specified, cultures were grown in BG11 medium at 25\u0026deg;C under continuous cool-white fluorescent light (15 \u0026mu;mol photons m\u003csup\u003e\u0026minus;2\u003c/sup\u003e s\u003csup\u003e\u0026minus;1\u003c/sup\u003e). Mutants of\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e were\u0026nbsp;maintained\u0026nbsp;in BG11 medium supplemented with 50 \u0026mu;g mL\u003csup\u003e\u0026minus;1\u003c/sup\u003e spectinomycin, and no antibiotics were added during the experimental treatments.\u003c/p\u003e\n\u003cp\u003eTo simulate the natural dehydration-rehydration cycle of \u003cem\u003eN. flagelliforme\u003c/em\u003e, exponentially growing cultures pre-adapted to nitrogen-replete or nitrogen-deplete medium were collected on\u0026nbsp;0.45 \u0026mu;m nitrocellulose membrane filters (Millipore), respectively. The samples were air-dried for 21 hours at 25\u0026deg;C, 70% relative humidity, and an irradiance of\u0026nbsp;15 \u0026mu;mol photons m\u003csup\u003e\u0026minus;2\u003c/sup\u003e s\u003csup\u003e\u0026minus;1\u003c/sup\u003e, followed by rehydration on\u0026nbsp;BG11 or BG11\u003csub\u003e0\u003c/sub\u003e (combined nitrogen-free) plates containing 1.5% agarose for 3 hours. This drying-rewetting cycle was repeated six times. Subsequently, the strains were recovered in liquid BG11 medium for assessment of cellular activity.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eRNA extraction and RT-qPCR analysis.\u0026nbsp;\u003c/strong\u003eTotal RNA was extracted from\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e using\u0026nbsp;Trizol Reagent (Cwbio, China)\u003csup\u003e61\u003c/sup\u003e. Genomic DNA was digested, and cDNA was synthesized using the\u0026nbsp;PrimeScript reagent gDNA Eraser Kit (Takara)\u0026nbsp;according to the manufacturer\u0026rsquo;s instructions. Transcript levels of target genes were quantified by\u0026nbsp;RT-qPCR\u0026nbsp;using a\u0026nbsp;7900HT Fast Real-Time PCR System (Thermo Fisher Scientific, USA)\u0026nbsp;with\u0026nbsp;SYBR Green Real-Time PCR Master Mix (Toyobo). The 16S rDNA gene was used as an internal reference gene. Relative expression levels were calculated using the\u0026nbsp;2\u003csup\u003e\u0026minus;\u0026Delta;\u0026Delta;CT\u003c/sup\u003e method. All primers used are listed in Supplementary Table 4.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eRNA preparation and\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eITC analysis.\u003c/strong\u003e The DNA fragment containing the\u0026nbsp;5\u0026prime; UTR of\u0026nbsp;\u003cem\u003ecgdA\u003c/em\u003e (designated Ctrl) was amplified using specific primers. Corresponding mutations\u0026nbsp;(M1, M2, and M3)\u0026nbsp;were then introduced into this fragment via fusion PCR. For \u003cem\u003ein vitro\u003c/em\u003e transcription, 20 \u0026mu;L reactions were prepared containing\u0026nbsp;50 mM Tris-HCl (pH 7.5), 15 mM MgCl\u003csub\u003e2\u003c/sub\u003e, 2 mM spermidine, 5 mM DTT, 2.5 mM of each\u0026nbsp;ribonucleoside 5\u0026prime; triphosphates\u0026nbsp;(NTP), and 20 units of\u0026nbsp;T7 RNA polymerase (Vazyme Biotech)\u003csup\u003e62\u003c/sup\u003e. Reactions were incubated at 37\u0026deg;C for 2 h, and the resulting RNA was purified using\u0026nbsp;the RNA Isolation Kit V2 (Vazyme Biotech)\u0026nbsp;according to the manufacturer\u0026rsquo;s instructions.\u003c/p\u003e\n\u003cp\u003eIsothermal titration calorimetry (ITC) experiments were conducted using a TA Instruments Nano ITC equipped with a low-volume sample cell at 25\u0026deg;C\u003csup\u003e28\u003c/sup\u003e. Guanidine hydrochloride (Sigma-Aldrich) was dissolved in Milli‑Q water (18.2 M\u0026Omega; cm\u003csup\u003e\u0026minus;1\u003c/sup\u003e) to a concentration of 1.4 mM and then titrated into the sample cell containing 0.2 mM RNA. A total of 25 serial injections of 2 \u0026mu;L each were performed, with 180‑second intervals between injections. Data were fitted to a one‑site binding model and analyzed using the Nanoanalyze software.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eLuciferase reporter assays.\u0026nbsp;\u003c/strong\u003eThe\u0026nbsp;luciferase\u0026nbsp;reporter system was constructed as described below. Using the\u0026nbsp;Ctrl-\u003cem\u003eluxAB\u003c/em\u003e strain as an example, the\u0026nbsp;5\u0026prime; UTR of \u003cem\u003ecgdA\u003c/em\u003e was fused\u0026nbsp;to the\u0026nbsp;\u003cem\u003eluxAB\u003c/em\u003e reporter fragment. The resulting DNA fragment was cloned into the\u0026nbsp;pRL25C\u0026nbsp;vector to generate the recombinant plasmid\u0026nbsp;pRL25C-Omega-\u003cem\u003ePromoter\u003c/em\u003e-\u003cem\u003eluxAB\u003c/em\u003e. This plasmid was introduced into\u0026nbsp;\u003cem\u003eN. flagelliforme\u0026nbsp;\u003c/em\u003eCCNUN1\u0026nbsp;using the conjugation method\u003csup\u003e63\u003c/sup\u003e. Transformants were selected on\u0026nbsp;1.5% agar BG11 plates containing 50 \u0026mu;g mL\u003csup\u003e\u0026minus;1\u003c/sup\u003e spectinomycin under 15 \u0026mu;mol photons m\u003csup\u003e\u0026minus;2\u003c/sup\u003e s\u003csup\u003e\u0026minus;1\u003c/sup\u003e illumination. Positive single colonies were verified by PCR and sequencing. Primers used are listed in Supplementary Table 4, and the resulting plasmids and strains are summarized in Supplementary Table 5.\u003c/p\u003e\n\u003cp\u003ePromoter activity was determined by measuring the relative bioluminescence intensity of luciferase in\u0026nbsp;\u003cem\u003eN. flagelliforme\u0026nbsp;\u003c/em\u003eCCNUN1 cells\u003csup\u003e61\u003c/sup\u003e. Aliquots of 0.2 mL culture, each containing an equal amount of chlorophyll \u003cem\u003ea\u003c/em\u003e, were transferred to a 96-well plate. Then, 5 \u0026mu;L of 0.1 mM decanal (dissolved in\u0026nbsp;dimethyl sulphoxide) was added as the substrate. Bioluminescence was measured at room temperature using a\u0026nbsp;BioTek Synergy 2 Plate Reader (BioTek). The recorded luminescence values were normalized to the corresponding\u0026nbsp;chlorophyll \u003cem\u003ea\u003c/em\u003e content.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMutant construction.\u003c/strong\u003e Chromosomal mutagenesis of the guanidine riboswitch in\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e was performed using\u0026nbsp;CRISPR-Cpf1\u0026nbsp;genome editing\u003csup\u003e32\u003c/sup\u003e. Briefly, the \u0026ldquo;TTN\u0026rdquo; motif within the guanidine‑I\u0026nbsp;aptamer\u0026nbsp;was identified, and a 20 bp DNA sequence downstream of this motif was inserted into the AarⅠ\u0026nbsp;site of the pCpf1 vector. Subsequently, the wild‑type fragment (Ctrl) and the mutated fragments (M2 and M3) were separately cloned into the BglⅡ\u0026nbsp;site of the same vector. The resulting plasmids were transformed into\u0026nbsp;\u003cem\u003eE. coli\u003c/em\u003e HB101\u0026nbsp;and then transferred into\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e via conjugation\u003csup\u003e63\u003c/sup\u003e. Mutants were selected on BG11 agar plates containing\u0026nbsp;50 \u0026mu;g mL\u003csup\u003e\u0026minus;1\u003c/sup\u003e spectinomycin under 15 \u0026micro;mol photons m\u003csup\u003e\u0026minus;2\u003c/sup\u003e s\u003csup\u003e\u0026minus;1\u003c/sup\u003e. Positive recombinant strains were verified using PCR and sequencing. All primers are listed in Supplementary Table 4, and the resulting plasmids and strains are summarized in Supplementary Table 5.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMetabolic analysis and isotope labeling experiments.\u0026nbsp;\u003c/strong\u003eThe\u0026nbsp;\u003cem\u003eN. flagelliforme\u003c/em\u003e strains\u0026nbsp;that had undergone periodic dehydration and rehydration were scraped from the filter membrane and quenched in cold methanol\u003csup\u003e64\u003c/sup\u003e. After centrifugation, the pellet was ground in liquid nitrogen, followed by three rounds of extraction with\u0026nbsp;500 \u0026mu;L of a 50:50 (v/v) methanol:water mixture. The supernatants from each extraction were pooled, dried in a vacuum centrifuge, and reconstituted in\u0026nbsp;500 \u0026mu;L Milli-Q water (18.2 M\u0026Omega; cm\u003csup\u003e\u0026minus;1\u003c/sup\u003e). The resulting extracts were filtered through a 0.22 \u0026mu;m nylon membrane filter (ANPEL, Shanghai, China) prior to analysis.\u003c/p\u003e\n\u003cp\u003eMetabolite extracts were analyzed using a\u0026nbsp;Shimadzu UFLC-30A\u0026nbsp;High-Performance Liquid Chromatograph\u0026nbsp;(HPLC) system coupled to an\u0026nbsp;Applied Biosystems 4500 QTRAP\u0026nbsp;mass spectrometer\u0026nbsp;(MS). Chromatographic separation was performed using a\u0026nbsp;Waters ACQUITY UPLC\u0026reg; BEH Amide column (2.1\u0026times;100 mm, 1.7 \u0026micro;m particle size). The mobile phase consisted of solvent A (water containing\u0026nbsp;10% ammonium hydroxide and 50 mM ammonium formate) and solvent B (acetonitrile). A linear gradient was applied as follows: solvent A was initially set at 10%, increased to 90% over 9 min, held for 1 min, returned to 10% within 0.5 min, and\u0026nbsp;maintained for 2.5 min. The injection volume was 4 \u0026micro;L, flow rate was\u0026nbsp;0.4 mL min\u003csup\u003e\u0026minus;1\u003c/sup\u003e, and column temperature was maintained at 40\u0026deg;C. Metabolites were detected in the positive ion mode using a quadrupole-linear ion trap mass spectrometer. Data were acquired using the\u0026nbsp;AB Sciex Analyst 1.6 software.\u003c/p\u003e\n\u003cp\u003eTo examine the potential presence of a guanidine carboxylation pathway, \u003cem\u003eN. flagelliforme\u003c/em\u003e was first pre‑adapted in nitrogen‑deplete medium and harvested onto 0.45 \u0026mu;m nitrocellulose membrane filters. The samples were air‑dried for dehydration and then rehydrated on a BG11\u003csub\u003e0\u003c/sub\u003e plate supplemented with 1 mM uniformly labeled U-[\u003csup\u003e13\u003c/sup\u003eC,\u003csup\u003e15\u003c/sup\u003eN]-arginine. Following 3 days of repeated dehydration\u0026ndash;rehydration cycles, the samples were collected for analysis. U-[\u003csup\u003e13\u003c/sup\u003eC,\u003csup\u003e15\u003c/sup\u003eN]-arginine was purchased from Cambridge Isotope Lab, and guanidine hydrochloride, arginine, and succinate were obtained from Sigma‑Aldrich. Standard solutions were prepared at defined concentrations, and a subset of the identified compounds was verified by matching their mass and retention time to those of authenticated standards.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eDetermination of photosynthetic activity.\u003c/strong\u003e Photosynthetic oxygen evolution was measured using a Clark-type oxygen electrode (Chlorolab 2, Hansatech Instruments)\u003csup\u003e33\u003c/sup\u003e. After periodic dehydration and rehydration, samples were scraped from the filters and resuspended in the reaction medium (BG11\u0026nbsp;medium\u0026nbsp;buffered with 25 mM Bis‑Tris propane, pH 8.0, and supplemented with 1 mM\u0026nbsp;NaHCO\u003csub\u003e3\u003c/sub\u003e). The electrode was calibrated with air‑equilibrated distilled water (maximum signal) and with a\u0026nbsp;Na\u003csub\u003e2\u003c/sub\u003eS\u003csub\u003e2\u003c/sub\u003eO\u003csub\u003e4\u003c/sub\u003e solution\u0026nbsp;(minimum signal). Temperature\u0026nbsp;was maintained at 25\u0026deg;C\u0026nbsp;using a\u0026nbsp;Polystat refrigerated bath (Cole-Parmer Instrument, Vernon Hills, IL). Net O\u003csub\u003e2\u003c/sub\u003e evolution was measured at\u0026nbsp;960 \u0026mu;mol photons m\u003csup\u003e\u0026minus;2\u003c/sup\u003e s\u003csup\u003e\u0026minus;1\u003c/sup\u003e, and respiration was measured as\u0026nbsp;O\u003csub\u003e2\u003c/sub\u003e consumption\u0026nbsp;in the dark.\u003c/p\u003e\n\u003cp\u003eChlorophyll fluorescence was measured with a\u0026nbsp;Plant Efficiency Analyser (Hansatech Instruments). Prior to measurement, samples were dark-adapted for 15 min to ensure that all PSII reaction centers were in the open state\u003csup\u003e33\u003c/sup\u003e. The maximal fluorescence (\u003cem\u003eF\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e) was determined by applying a pulse of saturating red light, and the maximum quantum yield of PSII (\u003cem\u003eF\u003c/em\u003e\u003csub\u003ev\u003c/sub\u003e/\u003cem\u003eF\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e) was calculated as\u0026nbsp;(\u003cem\u003eF\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e \u0026ndash;\u0026nbsp;\u003cem\u003eF\u003c/em\u003e\u003csub\u003eo\u003c/sub\u003e)/\u003cem\u003eF\u003c/em\u003e\u003csub\u003em\u003c/sub\u003e.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMeasurement of nitrogenase activity.\u0026nbsp;\u003c/strong\u003eNitrogenase\u0026nbsp;activity in\u0026nbsp;\u003cem\u003eN. flagelliforme\u0026nbsp;\u003c/em\u003estrains was determined using the acetylene reduction assay\u003csup\u003e65\u003c/sup\u003e. Following treatment, cells were scraped from the filter and resuspended in 2 mL of\u0026nbsp;BG11\u003csub\u003e0\u003c/sub\u003e medium\u0026nbsp;inside a 10‑mL sealed tube and adjusted to a final\u0026nbsp;chlorophyll\u0026nbsp;\u003cem\u003ea\u003c/em\u003e content of 0.1 mg. The headspace of each tube was repeatedly evacuated and replaced with\u0026nbsp;C\u003csub\u003e2\u003c/sub\u003eH\u003csub\u003e2\u003c/sub\u003e gas. Samples were then incubated at\u0026nbsp;5 \u0026mu;mol photons m\u003csup\u003e\u0026minus;2\u003c/sup\u003e s\u003csup\u003e\u0026minus;1\u003c/sup\u003e and 25\u0026deg;C\u0026nbsp;for 6 h, after which the gas phase was collected and analyzed on an\u0026nbsp;Agilent 7650A gas chromatograph. Data are presented as mean values from at least three replicate cultures.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eTranscriptomic analysis.\u0026nbsp;\u003c/strong\u003eTriplicate cultures of\u0026nbsp;\u003cem\u003ecgdA::\u003c/em\u003eCtrl and \u003cem\u003ecgdA::\u003c/em\u003eM2 strains\u0026nbsp;were collected after rehydration following dehydration and were immediately frozen in liquid nitrogen. Total RNA was extracted using\u0026nbsp;TRIzol Reagent according to the manufacturer\u0026rsquo;s instructions (Invitrogen). RNA concentration was measured using an\u0026nbsp;ND-2000 spectrophotometer (NanoDrop), and the samples were subsequently sent to\u0026nbsp;Majorbio Co. Ltd. for library preparation\u0026nbsp;and 150‑bp\u0026nbsp;paired-end sequencing on an Illumina HiSeq\u0026times;TEN\u0026nbsp;platform. Ribosomal RNA sequences were removed using\u0026nbsp;SortMeRNA v1.9\u003csup\u003e66\u003c/sup\u003e, and the resulting clean reads were aligned to the reference genome using\u0026nbsp;Rockhopper\u003csup\u003e67\u003c/sup\u003e. Normalization factors were calculated using the Trimmed Mean of M‑values method, and differential expression analysis was performed with\u0026nbsp;edgeR v3.20.7\u003csup\u003e68\u003c/sup\u003e. Gene expression profiles were further analyzed statistically using the Mfuzz package from Bioconductor in R\u003csup\u003e69\u003c/sup\u003e. A full list of the differentially expressed genes is available in\u0026nbsp;Supplementary Data 1.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eIn silico analysis of guanidine catabolism genes.\u003c/strong\u003e Homologs of CgdA, CgdB, GcA, AtzF, GdmH, GhaA, GhaB, ABC1-3, SssF, and Efe\u0026nbsp;were identified using BLAST searches against the NCBI databases. For phylogenetic analysis, two datasets were separately aligned using\u0026nbsp;the MAFFT multiple alignment program\u003csup\u003e70\u003c/sup\u003e: one comprising a concatenation of 31 single‑copy protein‑coding genes conserved across all representative species, and the other consisting of 16S rDNA sequences from representative cyanobacteria and bacteria. A maximum‑likelihood phylogenetic tree was constructed with\u0026nbsp;RAxML v8.1.20 under the PROTGAMMA model, with branch support evaluated by 1,000 bootstrap replicates, as previously described\u003csup\u003e10\u003c/sup\u003e. Basic genomic statistics for the 2,020 cyanobacterial genomes used in the comparative analysis are provided in Supplementary Table 2. The occurrence and frequency of the RNA motif RF00442 (guanidine‑I aptamer) were examined using\u0026nbsp;Infernal (v.1.1.2), with CMsearch applied to the motif at an\u0026nbsp;\u003cem\u003eE\u003c/em\u003e-value threshold of 0.0001\u0026nbsp;against all replicons from the included species\u003csup\u003e71\u003c/sup\u003e.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eStatistical analyses.\u003c/strong\u003e All experiments were performed with a minimum of three independent replicates. Statistical processing was conducted using Origin 2021\u0026nbsp;software\u0026nbsp;(OriginLab). For comparisons across multiple treatment groups, Tukey\u0026rsquo;s honest significant difference (HSD) test was applied, with a\u0026nbsp;\u003cem\u003eP\u003c/em\u003e-value\u0026lt;0.05\u0026nbsp;considered statistically significant. Differences between two independent groups were evaluated using the Student\u0026rsquo;s\u0026nbsp;\u003cem\u003et\u003c/em\u003e-test.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eData availability.\u0026nbsp;\u003c/strong\u003eData supporting the findings of this study are provided in the Supplementary Data files. Transcriptomic data have been deposited in the NCBI SRA database under accession code PRJNA1099419: https://www.ncbi.nlm.nih.gov/search/all/?term=PRJNA1099419. Guanidine-I aptamers and guanidine catabolism genes were identified using BLAST searches of NCBI databases, and the corresponding results are presented in Tables S2 and S3.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgement\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was funded by the National Natural Science Foundation of China (No. 32430005, No. 32270397, and No. 32470408), the China Postdoctoral Science Foundation (No. 2019T120675), and the Tianchi Talent Program of the Xinjiang Uygur Autonomous Region.\u003c/p\u003e\u003cp\u003e\u003cstrong\u003eAuthor contributions\u003c/strong\u003e: J.L.S., and B.S.Q. designed the research project; J.L.S., L.Z., Y.C., and Y.L. performed research; J.L.S., G.W.Q., L.Z., H.F.X., Z.C.Z., G.Z.D., S.H., C.Y., W.R.H., and B.S.Q. analyzed data; and J.L.S., G.W.Q., W.R.H., and B.S.Q. wrote the paper.\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interest.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eOren, N., Raanan, H., Murik, O., Keren, N. \u0026amp; Kaplan, A. 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Infernal 1.1: 100-fold faster RNA homology searches. \u003cem\u003eBioinformatics\u003c/em\u003e \u003cstrong\u003e29\u003c/strong\u003e, 2933\u0026ndash;2935 (2013).\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":true,"hideJournal":true,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"cyanobacteria, desert, desiccation tolerance, Guanidine, guanidine-I riboswitch, resurrection","lastPublishedDoi":"10.21203/rs.3.rs-8507520/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-8507520/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eIn desert ecosystems, microbial activity is driven by brief hydration pulses but severely constrained by persistent nutrient scarcity\u003csup\u003e1–4\u003c/sup\u003e. Cyanobacteria, as essential pioneer photoautotrophs, sustain biogeochemical cycling and ecosystem stability in these arid landscapes, yet their rapid resuscitation following rewetting is critically limited by nitrogen availability\u003csup\u003e3,4\u003c/sup\u003e. Although the nitrogen demand is met later by biological nitrogen fixation, it is irrelevant during early rehydration due to the high energy costs and delayed nitrogenase activation, creating a critical metabolic bottleneck\u003csup\u003e5–10\u003c/sup\u003e. Here we demonstrate that the desert cyanobacterium \u003cem\u003eNostoc flagelliforme \u003c/em\u003eovercomes this constraint by activating a previously overlooked guanidine carboxylase pathway, which sustains the rapid remobilization of internal nitrogen reserves upon rehydration. Pathway activity correlates strongly with hydration and nitrogen status. Genetic experiments demonstrate its essential role in recovery from desiccation and characterize the key role of a guanidine-I riboswitch. Phylogenetic evidence indicates that pathway genes are transferred horizontally, correlating with habitat aridity. Our findings establish a novel ecological role for guanidine in desert ecosystems, and suggest its impact on microbial community assembly in nutrient-poor drylands. This work reveals a conserved adaptation mechanism that supports cyanobacterial resilience in xeric environments and provides a physiological basis for enhancing biocrust restoration strategies.\u003c/p\u003e","manuscriptTitle":"Guanidine fuels rapid resurrection of desert cyanobacteria","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-01-22 08:58:52","doi":"10.21203/rs.3.rs-8507520/v1","editorialEvents":[],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"47d967dc-f3a2-42dc-b96b-c1599bb89e9a","owner":[],"postedDate":"January 22nd, 2026","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[{"id":60623312,"name":"Biological sciences/Microbiology/Environmental microbiology/Soil microbiology"},{"id":60623313,"name":"Earth and environmental sciences/Ecology/Ecophysiology"}],"tags":[],"updatedAt":"2026-01-22T15:06:00+00:00","versionOfRecord":[],"versionCreatedAt":"2026-01-22 08:58:52","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-8507520","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-8507520","identity":"rs-8507520","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}