Unraveling the diversity and functional potential of cyanosphere microbiomes assembled from terrestrial cyanobacteria

Unraveling the diversity and functional potential of cyanosphere microbiomes assembled from terrestrial cyanobacteria | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (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];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Unraveling the diversity and functional potential of cyanosphere microbiomes assembled from terrestrial cyanobacteria View ORCID Profile Brianne Palmer , View ORCID Profile Estelle M Couradeau , Jeffrey R. Johansen , View ORCID Profile Tania Kurbessoian , View ORCID Profile Jose Ortega Carranza , View ORCID Profile Jason E Stajich , Ryan Ward , View ORCID Profile Nicole Pietrasiak doi: https://doi.org/10.1101/2025.06.10.658773 Brianne Palmer 1 Bonn Institute of Organismic Biology, University of Bonn , Bonn, Germany Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Brianne Palmer For correspondence: bpalmer{at}uni-bonn.de nicole.pietrasiak{at}unlv.edu Estelle M Couradeau 2 Department of Ecosystem Science and Management, The Pennsylvania State University, University Park , PA, USA 3 The Huck Institutes of the Life Sciences, The Pennsylvania State University, University Park , PA, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Estelle M Couradeau Jeffrey R. Johansen 4 Biology Department, John Carroll University, University Heights , OH, USA 5 Faculty of Sciences, Botany Department, University of South Bohemia , České Budějovice, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tania Kurbessoian 6 Department of Microbiology and Plant Pathology, University of California-Riverside , Riverside, CA USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Tania Kurbessoian Jose Ortega Carranza 7 Plant and Environmental Sciences Department, New Mexico State University , Las Cruces, NM, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jose Ortega Carranza Jason E Stajich 6 Department of Microbiology and Plant Pathology, University of California-Riverside , Riverside, CA USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jason E Stajich Ryan Ward 7 Plant and Environmental Sciences Department, New Mexico State University , Las Cruces, NM, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Nicole Pietrasiak 7 Plant and Environmental Sciences Department, New Mexico State University , Las Cruces, NM, USA 8 School of Life Sciences, University of Nevada - Las Vegas , Las Vegas, NV, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Nicole Pietrasiak For correspondence: bpalmer{at}uni-bonn.de nicole.pietrasiak{at}unlv.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract The cyanosphere consists of heterotrophic microorganisms residing within the exopolysaccharide sheath of cyanobacteria, acting as a crucial interface between the cyanobacteria and their surrounding environment. Understanding the interactions between cyanobacteria and their cyanospheres is essential for predicting the success of terrestrial cyanobacteria in providing ecosystem services in nutrient-poor environments. However, knowledge of the microbial diversity within the cyanosphere remains limited. In this study, we employed metagenomic sequencing to reconstruct 410 metagenome-assembled genomes (MAGs) from cyanosphere-associated microbes linked to 56 unialgal terrestrial cyanobacteria cultures, representing 12 distinct cyanobacteria orders. Our findings revealed that the composition of cyanosphere microbial communities was unique to each cyanobacterial host and was significantly shaped by environmental factors such as habitat, precipitation, and temperature from which the cultures were originally obtained. Notably, three microbial genera, Brevundimonas , Devosia , and Sphingopyxis , were present in over 30% of the cyanospheres, forming a core cyanosphere microbiome. Functional gene analysis showed a distinction between the cyanobacteria and their associated cyanospheres, with dissimilatory nitrate reduction being the dominant pathway in the cyanosphere, while nitrogen fixation was more common in the cyanobacteria. Three cyanospheres also contained nitrogen fixation genes of which two hosts were nitrogen fixation capable themselves. The cyanosphere harbored genes for polysaccharide lyases, indicating a possible link to the exopolysaccharides produced by the cyanobacteria. Given the observed variability in microbial community composition and function across different cyanobacterial hosts, future ecological assessments and restoration efforts involving cyanobacteria should not only focus on the cyanobacteria themselves but also consider their associated microbial communities. Importance Our study identifies members of a highly understudied, and potentially under-valued, microbial community -- the cyanosphere. We used a diversity of terrestrial cyanobacteria to understand how the cyanosphere composition and predicted functions were influenced by the host cyanobacterium and environmental factors using metagenomics. This is a new approach to study the cyanosphere and provides insights into the diversity of terrestrial microbial communities. Importantly, our results underscore the need to consider microbial consortia when assessing the ecological potential of cyanobacteria in terrestrial restoration. Introduction Cyanobacteria are prevalent and perform various ecosystem functions in almost any environment where sunlight is at least temporarily available. This includes terrestrial environments which are highly complex and are characterized by often very contrasting abiotic and biotic conditions such as found in soil, hydroterrestrial, and subaerial habitats ( 1 , 2 ). In soils, cyanobacteria contribute to carbon sequestration via organic and inorganic pathways ( 3 ), and enrich soil fertility by fixing atmospheric nitrogen and reducing soil erosion ( 4 , 5 ). Cyanobacteria are also foundational components of biological soil crusts which cover approximately 30 % of drylands globally ( 6 ). In hydroterrestrial environments, such as periodically dry vernal pools, algal and cyanobacterial biofilms often form rapidly after rehydration and are important for maintaining local food webs as microscopic primary producers ( 7 , 8 ). In subaerial habitats, which include exposed surfaces like rocks and tree bark, cyanobacteria are pioneers in colonizing these environments, contributing to the formation of epilithic and epiphytic biofilms that support other microbial and plant life ( 9 – 11 ). As early colonizers on these substrates, the cyanobacteria can enrich the substrate with organic carbon through photosynthesis and can improve nutrient cycling in the micro-environment by trapping dust and soil particles and fixing atmospheric nitrogen ( 10 , 12 ). A distinctive feature of many cyanobacteria is the presence of the cyanosphere ( 13 ). This is an ecological niche akin to the rhizosphere of plants. The cyanosphere is a microenvironment surrounding cyanobacterial cells, especially in filamentous forms, and is inhabited by heterotrophic microorganisms that closely associate with their cyanobacterial hosts ( 13 ). These heterotrophic microorganisms may benefit from the organic compounds produced by the cyanobacteria through photosynthesis, and in return, they can assist the cyanobacteria by providing essential nutrients ( 14 ). Mutualistic interactions are common in the rhizosphere and may thus also occur within the cyanosphere ( 15 , 16 ). Current research on the terrestrial cyanosphere is limited. Previous work on the cyanosphere has focused on Microcoleus vaginatus , a filamentous cyanobacteria known to be an early colonizer of biological soil crusts without the ability to fix nitrogen. In one of the first cyanosphere studies, Couradeau and her colleagues found that most of the inhabitants of the M. vaginatus cyanosphere were copiotrophs, and some were diazotrophs and contained more nitrogen-fixation genes in comparison to the bulk soil microbiome. This highlights the potential role of nutrient exchange of carbon and nitrogen between the cyanobacteria and the diazotrophs in the cyanosphere. A subsequent study found evidence of a carbon-nitrogen exchange between M. vaginatus and the cyanosphere ( 14 ). The success of M. vaginatus as an early colonizer and important species for cyanobacterial-mediated dryland soil restoration may be because of the heterotrophs within the cyanosphere such as Arthrobacter, Massillia, and Bacillus which, when added to the cyanobacterial growth medium, increased cyanobacterial growth ( 17 ). Nelson and his colleagues found evidence of signaling between M. vaginatus and its cyanosphere, further emphasizing the interconnectedness between the cyanobacteria and their associated heterotrophs ( 18 ). Beyond M. vaginatus, a few studies have assessed the microbial community composition within the cyanosphere with most studies focused on either soil or freshwater habitats. In a direct comparison of the rhizosphere and the cyanosphere. Another study examined growth-promoting strains in both the rhizosphere and biocrust cyanospheres. They found 18 phyla common to both and identified growth-promoting isolates such as Bosea and Pseudoarthrobacter in the cyanosphere ( 16 ). Within aquatic environments, one study used metatranscriptomics to study the cyanospheres of both nitrogen-fixing and non-nitrogen-fixing aquatic cyanobacteria Dolichospermum and Microcystis showing that the bacterial associations were determined by the cyanobacterial host with the most abundant phyla being Bacteroidetes, Gemmatimonadetes, and Proteobacteria. They also identified functional redundancy within the cyanosphere ( 19 ). However, the diversity and ecology of hydroterrestrial and subaerial habitats are virtually unexplored. Based on the limited research of terrestrial cyanospheres, we know that the identity of the microbes within the cyanosphere can be important for the success of the cyanobacteria ( 17 ), and the functions of the cyanosphere microbes can provide essential nutrients to the cyanobacteria ( 13 , 14 , 18 ). However, the breadth of terrestrial cyanobacterial species is large, and current research is limited to M. vaginatus. Therefore, there is a need to understand the identity and function of the cyanosphere microbes within the cyanosphere from a diverse set of terrestrial cyanobacteria. The use of next-generation sequencing techniques allows for the identification of the microbial community within the cyanosphere. Metagenomics, in particular, provides information about the identity of the microbes and the potential functions they may be performing. Here, we used metagenomic sequencing on 56 cultured terrestrial cyanobacteria and their cyanosphere microbiomes. These cyanobacteria are classified into 12 of the currently recognized 19 orders in cyanobacteria and represent broad phylogenetic diversity. We sought to understand how the taxonomy of the host cyanobacteria and the environment affects the heterotrophic community of the cyanosphere. Methods Cyanobacteria cultures Metagenomes were obtained from unialgal polycultures containing cyanobacteria and associated heterotrophic cyanosphere microbiomes. Polycultures were established via traditional dilution plating of environmental samples obtained from various collection trips led by Johansen, Pietrasiak, and collaborators. Collection years and detailed information can be found in Ward et al. (2021). Polycultures were isolated between 1965 and 2015 and represent reference cultures of recently well-studied taxonomic and phylogenetic benchmarks in cyanobacteria taxonomy and systematics. The dominant cyanobacterial members in these cultures are phylogenetically diverse, spanning 12 orders of cyanobacteria. Cultures have been maintained in two culture collections at John Carroll University (JCU) and University of Nevada - Las Vegas (UNLV) on solid or liquid Z8 media, an oligotrophic freshwater algal medium ( 20 ). Although dates of isolation span 50 years, close to identical culture conditions (use of Z8 medium, 16/8 hr light/dark photoperiod, low light intensity, grown in climate-controlled chambers) have been maintained at JCU since 1992 and have been reproduced at UNLV. At the time of DNA extraction, the cultures’ microbial biomass was dominated by cyanobacteria. Metagenomic Sequencing Polyculture biomass was grown in liquid Z8 media, harvested, flash-frozen, and stored before DNA extraction using the Qiagen DNeasy PowerLyzer kit with bead-beating, following detailed protocols available on protocols.io (dx.doi.org/10.17504/protocols.io.brg4m3yw) ( 2 ). Extracted DNA was stored at -20°C prior to shipment to the Joint Genome Institute (JGI). JGI performed the library preparation and next gen Illumina sequencing steps as follows. DNA library preparation was performed using the KAPA Biosystems high-throughput library preparation kit p/n KK8235 on a PerkinElmer Sciclone next-generation sequencing (NGS) robotic liquid handling system. Then, 200 ng of sample DNA was sheared to 300 bp using a Covaris LE220 focused ultrasonicator. The sheared DNA fragments were size selected by double solid-phase reversible immobilization (SPRI). The selected fragments were end repaired, A-tailed, and ligated with Illumina-compatible sequencing adaptors from IDT containing a unique molecular index barcode for each sample library. The prepared libraries were quantified with a KAPA Biosystems quantitative PCR (qPCR) kit on a Roche LightCycler 480 real-time PCR instrument. Genomic libraries were sequenced with a NovaSeq instrument (Illumina, San Diego, CA) using NovaSeq XP V1 reagent kits and an S4 flow cell following a 2 x 150-bp indexed run recipe. Demultiplexed reads were processed with BBDuk v38.87 ( 17 ) to remove contaminants, trim adapter sequence and “G” homopolymers ≥5 in size at the ends, quality trim reads, and remove reads with ≥4 “N” bases, with an average quality score of >3, or with a length of <51 bp. Raw sequence products were submitted to NCBI SRA archive. A list of BioProjects is provided in Ward et al. ( 2 ) with additional descriptions in Table S1. Microbial community analyses Sequence Read Archive (SRA) reads were first uploaded into the KBase platform ( 21 ). The quality of these reads was assessed using the Fast-QC tool ( 22 ), and those that did not meet the quality threshold (Q score > 30) were refined using Trimmomatic ( 23 ). The reads for each Cyanobacteria were assembled using metaSPAdes ( 24 ). The assembled contigs were then segregated into Metagenome-Assembled-Genomes (MAGs) via MaxBin2 ( 25 ). Taxonomic classification was conducted using GTDB-tk ( 26 ). A comprehensive taxonomy table, incorporating DRAM annotation, was generated to establish a connection between taxonomy and function. MAGs that did not meet the minimum requirements for medium-quality MAGs (> 50% completion and < 10% contamination) ( 27 ) were removed. The KBase narrative is publicly available at https://narrative.kbase.us/narrative/202318 . All subsequent analyses were performed in R (version 4.3.1). The amp_alpha_diversity function within the R package ampvis2 was used to calculate alpha diversity metrics including the total number of MAGs and Shannon diversity ( 28 ). Differences in MAG richness and Shannon diversity between categorical variables were calculated using ANOVAs with Tukey HSD post-hoc tests. To determine if there is a core cyanosphere, we calculated the number of cyanospheres in which heterotrophic genus occurred. Genera found in > 30% of the cyanospheres were classified as part of the core cyanosphere. The package microeco was also used to create a heatmap of the most abundant genera. Bray-Curtis distances were used to calculate the beta diversity using the microeco package. To assess differences in the microbial community composition across categorical variables, a PERMANOVA analysis was performed using the cal_manova function. Beta diversity was visualized through Principal Coordinates Analysis (PCoA) plots generated in microeco R package ( 29 ). Climate variables were obtained from BIOCLIM ( 30 ). When latitudinal and longitudinal data were available, we identified the mean annual temperature and the annual precipitation at the collection location of each cyanobacterium in BIOCLIM. Due to the covariance between environmental variables, only mean annual temperature and precipitation were used. The impact of continuous variables such as temperature, precipitation, and altitude on microbial community composition was evaluated using the cal_ordination_envfit function within microeco . This relationship was further illustrated with a distance-based Redundancy Analysis (dbRDA) plot using the microeco R package ( 29 ). The functional abundances, as assigned via DRAM, were visualized using heat maps created with the pheatmap R package ( 31 ) for the cyanosphere community and the host Cyanobacteria. The code for this project is publicly available on GitHub at https://github.com/briannepalmer/Cyanosphere . Results Spatial representation Cyanobacterial hosts represented 14 locations spanning 4 continents (including the Pacific Islands) ( Figure 1 ). Soil Cyanobacteria were collected from 5 types of parent material (dolomite, granitic, gypsiferous, gneiss, and limestone) (Table S1). The altitude of the host Cyanobacteria ranged from 11 to 2043 meters. The annual precipitation was between 3 mm and 3229 mm. The annual mean temperature ranged from 5.2 °C to 23.6 °C. In total, 410 cyanosphere-associated microbial MAGs were identified, 214 from soil cyanobacteria, 159 from subaerial cyanobacteria, 23 from freshwater cyanobacteria, and 13 from hydroterrestrial cyanobacteria. Download figure Open in new tab Figure 1: Map of the locations where the cyanobacteria were collected. Five cyanobacteria ( Drouetiella lurida, Oculatella coburnii, Pleurocapsa ergoicii, Roholtiella mojaviensis and Timaviella radians ) did not have coordinate data and are not included on the map. The points are colored by the habitat where the cyanobacteria were collected, and the numbers are assigned to each cyanobacterium found in Table S1. Core and Most Abundant Genera Proteobacteria (72%), Bacteroidota (11%), and Actinobacteria (6%) were the most abundant phyla in the cyanospheres ( Figure 2A ). Within Proteobacteria, Rhizobiales was the most abundant order, occurring in 79% of the cyanospheres ( Figure 2B ). Only three genera were found in > 30% of the samples: Brevundimonas (40%) , Devosia (36%), and Sphingopyxis (30%) ( Figure 3 ). These genera could represent a core terrestrial cyanobacterial cyanosphere. There were two families in > 50% of the cyanospheres: Sphingomonadaceae and Caulobacteraceae. No genera were found in every cyanobacterial order. However, two genera – Allorhizobium and Sphingopyxis – were found in all four habitat types. Additionally, Brevundimonas, Devoisa, and Sphingomonas were found in the three terrestrial habitats (soil, subaerial, hydro-terrestrial). Download figure Open in new tab Figure 2: Bar plots depicting the relative abundance of the most abundant phyla (A) and the Proteobacteria orders (B) for the MAGs in the cyanosphere. Download figure Open in new tab Figure 3: Heatmap showing the relative abundance of the 50 most abundant heterotrophic genera found within each cyanobacterial host. Dark red indicates higher relative abundance while light red and white indicates low or no abundance in the given cyanosphere. The heterotrophic genera on the y-axis are ordered by the total relative abundance across all cyanospheres (x-axis). Alpha Diversity Metrics were Consistent Across Cyanobacterial Hosts MAG richness and Shannon diversity did not vary among cyanobacterial orders (ANOVA, P = 0.18, P = 0.06) or among habitats (ANOVA, P = 0.61, P = 0.54) ( Figure 4 ). Chroococcus sp., Trichotorquatus andrei, and Mojavia pulchra each had the most heterotrophic MAGs (n = 14) followed by Kastovskya adunca, Oscillatoria tanganyikae, and Microcoleus vaginatus (n = 13). Pelatocladus maninholoensis did not contain any non-cyanobacterial MAGs. Plectolyngbya sp., Trichormus sp., Scytonematopsis contorta, and Aphanocapsa sp. only contained 1 non-cyanobacterial MAG. There was an average of 8 MAGs per cyanobacteria. Download figure Open in new tab Figure 4: Bar charts depicting the total number of MAGs (A) and the Shannon diversity of the MAGs (B) for each Cyanosphere, grouped by habitat. The bars are colored by the order of the host Cyanobacteria. Microbial Community Composition Varies Between Habitats, Rather than Between Cyanobacterial Hosts Based on the PcOA plot and PERMANOVA (P = 0.08), there are no differences in the heterotrophic community composition among cyanobacterial host orders. The PCoA axis explain 12.7% and 9.6% of the variation in the community composition and there is little clustering between heterotrophic communities with the same cyanobacterial host order ( Figure 5A ). However, the microbial community composition did vary by host genera (PERMANOVA, P = 0.02) highlighting that each cyanosphere contains a unique microbial community. There was also a significant effect of habitat on the community composition (PERMANOVA, P = 0.001). The mean annual temperature (P < 0.001), annual precipitation (P < 0.001), and altitude (P 0.015) where the cyanobacteria were collected also influenced the community composition ( Figure 5B ). When the environmental factors and habitat type are included in an RDA plot, there is distinct grouping between the soil and subaerial samples and the axes explain 46.8% and 31.6% of the variation in the microbial community composition. The soil communities are grouped with the annual mean temperature vector, while the subaerial samples group with annual precipitation. Download figure Open in new tab Figure 5: Ordination plots showing the community composition based on the MAG genera in the cyanospheres. A) a PCoA depicting the MAG community composition colored by the order of the cyanobacterial host. B) a RDA plot colored by habitat indicating the grouping of the cyanosphere community composition. The ellipses show groupings of the cyanospheres with more than 3 representatives. The arrow vectors (Annual Precipitation, Annual Mean Temperature, and Altitude) show environmental variables that were significantly (P<0.05) correlated with the cyanosphere community composition. Cyanosphere and cyanobacterial hosts differ in key functions Only three cyanospheres contained cyanosphere members with nitrogen-fixation genes: Hassallia sp., Brasilonema angustatum, and Oscillatoria princeps ( Figure 6 ). These putative diazotrophs included Bradyrhizobium in Brasilonema angustatum and Oscillatoria princeps , as well as Skermanella in Hassallia sp. Download figure Open in new tab Figure 6: Heatmaps depicting the abundance of genes relating to nitrogen pathways for the cyanosphere (A) and cyanobacteria (B). The x-axis labels describe the nitrogen pathways including assimilatory nitrate reduction, complete nitrification, denitrification, dissimilatory nitrate reduction, nitrate assimilation, nitrification, nitrogen fixation, and nitrite + ammonia => nitrogen. Dissimilatory nitrate reduction emerged as the most prevalent nitrogen pathway within the cyanosphere ( Figure 6 ), although it was notably absent in the cyanospheres of Aphanothece saxicola and Plectolyngbya sp. The cyanospheres of Kastovskya adunca and Nostoc desertorum had the highest number of genes coding for dissimilatory nitrate reduction (DNRA), while denitrification was most prominent in the cyanosphere of Oscillatoria princeps . Genes for assimilatory nitrate reduction were most abundant in the cyanospheres of Nostoc desertorum and Mojavia pulchra , whereas Desmonostoc vinosum exhibited the most complete nitrification pathways. Nitrate assimilation was most pronounced in cyanosphere microbes of Kastovskya adunca , Nostoc desertorum , and Rexia erecta . Notably, none of the cyanospheres contained pathways for nitrification. This pattern contrasted with the nitrogen pathways observed in the cyanobacteria themselves. Myxaxorys chilensis , Plectolyngbya sp., and Trichocoleus desertorum lacked high-quality cyanobacterial MAGs, while Timaviella obliquedivisa had two cyanobacterial MAGs. Iphinoe sp. contained the highest number of genes for nitrogen fixation, followed by Brasilonema octagenarum and Brasilonema angustatum ( Figure 6 ). In total, 25 cyanobacterial species possessed nitrogen fixation genes, most corresponding with the order Nostocales. Both Hassallia sp. and Brasilonema angustatum , whose cyanosphere microbes had nitrogen fixation genes present contained cyanobacterial nitrogen fixation genes corresponding to their ability to produce heterocytes and thus taxonomic affiliation to the order Nostocales taxa, while Oscillatoria princeps , order Oscillatoriales, did not contain nitrogen fixation genes. Interestingly, cyanobacteria taxonomically not affiliating with the order Nostocales also contained nitrogen fixation encoding genes, such as, Pegethrix bostrychoides . As with the cyanospheres, no genes involved in nitrification were detected in the cyanobacteria. Carbohydrate-Active enZYme (CAZy) pathways were also prevalent in the cyanospheres, particularly those involving glycosyl transferases and glycoside hydrolases ( Figure 7 ). The cyanospheres of Chroococcus sp. and Roholtiella mojaviensis contained the highest number of genes related to glycosyl transferases, while Aphanocapsa sp. had the fewest. For glycoside hydrolases, Microcoleus vaginatus and Chroococcus sp. cyanospheres had the most genes, whereas Scytonematopsis contorta had the fewest. Although polysaccharide lyases were less abundant, they were still detected in 48 cyanospheres, with genes for polysaccharide lyases present in 68 genera ( Figure 7 ). The Nostoc desertorum cyanosphere contained the most polysaccharide lyase genes, while Scytolyngbya sp. had only one. Download figure Open in new tab Figure 7: Heatmaps depicting the abundance of genes relating to CAZy pathways found in the cyanosphere (A) and the cyanobacteria (B). The genes present in the cyanosphere differ notably from those found in the cyanobacterial host. In the cyanobacteria, Scytonematopsis contorta had a high abundance of glycosyl transferase genes, while Timaviella obliquedivisa possessed the most glycoside hydrolase-related genes ( Figure 7 ). Polysaccharide lyases, however, were relatively scarce in cyanobacteria, with only 24 species containing these genes. Of those, only Nostoc indistinguendum , Nostoc desertorum , Pleurocapsa ercegoicii , Roholtiella mojaviensis , and Scytolyngbya sp. had more than one gene associated with polysaccharide lyases. Interestingly, genes for carbohydrate-binding molecules and carbohydrate esterases were absent from all cyanospheres. Among the polysaccharide lyases identified, five alginate lyases were found—PL5, PL6, PL15, PL17—all capable of degrading alginate. Thirty cyanospheres, mostly from Nostocales (n=14), contained alginate lyases, while Synechococcales cyanospheres lacked these enzymes ( Figure 8 ). The Iphinoe sp. cyanosphere had the highest number of alginate lyases, followed by Microcoleus vaginatus . Download figure Open in new tab Figure 8: The total number of alginate lyases found in the cyanosphere of each cyanobacteria host. The colors represent the order of the cyanobacterial host. Discussion Using metagenomic sequencing, we identified non-cyanobacterial members of the cyanosphere from various habitats and cyanobacterial orders. This study is the first, to our knowledge, to identify the cyanosphere for a broad scope of terrestrial cyanobacteria and has implications for understanding interspecific interactions and the growth conditions necessary for cyanobacteria. Trends within cyanospheres Based on 410 non-cyanobacteria heterotrophs from 56 cyanobacterial hosts, we identified patterns between the cyanospheres. The most common genera, found in over 30% of the cyanospheres were Brevundimonas, Devosia, and Sphingopyxis. These genera might indicate a “core microbiome” as prior studies have used abundance in 30% of samples as an arbitrary cutoff for calculating core communities ( 32 , 33 ). Brevundimonas is a diverse genus and species have been found in both soil and aquatic environments ( 34 – 36 ). In the present study, Brevundimonas were found in soil, subaerial, and hydroterrestial environments and accounted for 6.8% of the MAGs in the cyanosphere. Brevundimonas species can live as free-living species in a variety of environments including as a diazotrophic plant-root colonizer ( 37 ), the aerobic and anaerobic conditions of activated sludge ( 38 ), alkaline soils ( 39 ), and thermal baths ( 40 ). The ability to thrive in such variable environmental conditions may be beneficial in the cyanosphere, as the cyanobacteria were collected from diverse environments. Devosia is a common genus in soil and aquatic sediment ( 41 – 45 ). It was also relatively common in the cyanospheres, found in 36% of the cyanospheres and representing 5.6% of the total MAGs. At least one species of Devosia can produce nitrogen-fixing root-nodule symbiosis with plants ( 46 ), indicating nitrogen fixation capability, which may also occur in the cyanosphere when nitrogen fixation is needed. However, there were no nitrogen-fixation genes detected in the cyanosphere Devosia in the present study. Two species of Devosia were previously isolated from the phycosphere of marine red alga ( 47 ). Previous work highlighted the genomic plasticity of Devosia , which allows this genus to utilize a wide variety of substrates ( 48 ). This ability to adapt to different environments may contribute to the prevalence of Devosia in the cyanosphere. In the cyanosphere, Sphingopyxis was found in 30% of the host cyanobacteria and represented 4.4% of the total cyanosphere MAGs. Like Devosia, Sphingopyxis is a widespread genus due to its ability to thrive in diverse and often stressful environments such as aquatic sediment ( 49 ), contaminated soil ( 50 , 51 ), and activated sludge ( 52 ). Likely contributing to its survival in stressful environments is the ability of several species to degrade environmental contaminants and toxins and produce secondary metabolites ( 53 ). The order, Rhizobiales was present in 79% of the cyanospheres, indicating that this order is an important constituent. Rhizobiales contains several nitrogen-fixing genera, primarily associated with root nodules, like Rhizobium. This is a broad taxonomic group, and not all genera can fix nitrogen. Interestingly, several Rhizobiales known to fix nitrogen were found in the cyanosphere, despite living in the cyanosphere of a nitrogen-fixing cyanobacteria including: Bosea, Mesorhizobium, Devosia, Bradyrhizobium, Methylobacterium, Allorhizobium, Microvirga, Ensifer, Neorhizobium, Agrobacterium, Shinella, and Pararhizobium ( 54 , 55 ). These genera are known root-nodule formers. However, we did not detect nif genes in their MAGs and currently we do not know if these microbes have an established symbiotic relationship with the cyanobacteria for carbon and nitrogen exchange like in plant roots. Rhizobiales that do not fix nitrogen were also found in these cyanospheres. These may be pathogenic, similar to the Rhizobiales root pathogens found in the rhizosphere ( 56 ). In the present study, only three cyanosphere microbes had nitrogen-fixing genes. This indicates that perhaps in most cases, the cyanosphere selects for members without the ability to fix nitrogen, or the metagenomes do not fully represent the metabolic diversity of the cyanosphere. The richness and diversity of the microbial community did not vary significantly between the cyanospheres while composition was influenced by the cyanobacterial host. A previous study with the rhizosphere showed that bacterial richness and diversity were greater in the bulk soil compared to the rhizosphere and that the richness and diversity of the rhizosphere rarely varied between rhizospheres ( 57 ). The authors state that the rhizosphere microbiota have similar traits regardless of the plant, thus resulting in similar richness and diversity measurements regardless of the phylogenetic placement of the bacteria. A similar phenomenon may occur in the cyanosphere, whereby bacteria filling particular niches colonize the cyanosphere regardless of cyanobacterial host phylogeny resulting in similar measurements of richness and alpha diversity between cyanospheres. Habitat influences cyanosphere microbiome The community composition of the cyanosphere was influenced more by the habitat where the cyanobacteria were collected than the phylogeny of the cyanobacteria. This suggests that the microbes forming the cyanosphere are not determined by relationships between the cyanosphere and the phylogenetic affiliation of the host cyanobacteria at the broader taxonomic level. The effect of habitat could be due to recruitment of the cyanosphere microbiome from the surrounding microbial community. This commonly occurs within the rhizosphere where the rhizosphere microbiome is a specific subset of the bulk soil microbiome ( 58 ). In the rhizosphere, root traits can explain the community composition ( 59 ). Thus, in the cyanosphere, the traits of the cyanobacteria could be the factors that determine which bacteria colonize the cyanosphere. One of the limits of this dataset is the lack of even sampling across locations. This should be rectified in future studies. As such, there was a bias between habitat and sampling location. For example, 21 of the 29 soil cyanospheres were collected from North America (primarily Mojave and Sonoran Deserts, Southern California). Thus, it is difficult to parse out if the difference in the community composition between habitats is due to the effect of the particular habitat or due to the effect of sampling location, in this case, with a bias towards microbial communities that are present in California. The problem is not persistent within the other habitat types as the subaerial samples ( 22 ) have 3 from Europe, 10 from North America, and 9 from the Pacific Islands and the freshwater samples are from Europe, North America, and the Pacific Islands. Furthermore, there was not even sampling across habitats with 29 samples from the soil, 22 subaerial, 3 fresh water, and 2 hydroterrestrial (both from Hawaii). This should not discount the results presented in our exploratory study thus far but rather should be used as a stepping stone for subsequent research. Given the differences in the microbial community composition between habitats (or locations), there is a need for further research to determine the specific factors that influence the cyanosphere microbial community composition. Cyanospheres and cyanobacteria differ in predicted nitrogen and carbon exchange pathways Unlike the rhizosphere, there is not a clear relationship of carbon for nitrogen exchange between the cyanosphere and the host cyanobacteria, perhaps because many cyanobacteria can fix nitrogen on their own, including 25 Cyanobacteria included in this study. However, three cyanospheres contained nitrogen-fixing genes within the cyanosphere: Hassallia sp., Brasilonema angustatum, and Oscillatoria princeps. Brasilonema angustatum and Hassallia sp., both of the order Nostocales, contain nitrogen-fixation genes in their genomes, are able to form heterocytes, and can fix atmospheric nitrogen ( 60 , 61 ). Thus, these cyanobacteria would not necessarily need to interact with nitrogen-fixers in their cyanosphere. Thus, it is puzzling why we found diazotrophic Bradyrhizobium and Skermanella in the cyanosphere of these two diazotrophic cyanobacteria. Bradyrhizobium is a common genus in the rhizosphere and can form root nodules ( 62 , 63 ), and both Skermanella and Bradyrhizobium were identified as key indicator species of potential diazotroph activity in terrestrial ecosystems ( 64 ). Perhaps under certain environmental conditions either the cyanobacteria or the cyanosphere bacteria ( Bradyrhizobium and Skermanella ) are not performing nitrogen fixation and gaining nitrogen through the symbiotic relationship. In comparison, Oscillatoria princeps did not have genes for nitrogen fixation, does not form heterocytes but recorded a Bradyrhizobium MAG in its cyanosphere, suggesting a possible mutualistic relationship with Bradyrhizobium in the cyanosphere. Neither the Cyanobacteria nor the cyanospheres had genes for nitrification. The nitrogen metabolism genes in the Cyanobacteria emphasize their role in assimilatory nitrate reduction and nitrogen fixation; while the cyanosphere has more genes related to dissimilatory nitrogen reduction and denitrification. Dissimilatory nitrate reduction reduces nitrate to nitrite through denitrification then reduces nitrite to ammonium, conserving nitrogen as ammonium rather than dinitrogen gas. Assimilatory nitrate reduction reduces nitrate to nitrite and ammonium which can be incorporated into the organism. The prevalence of these two different pathways of nitrogen reduction indicates the different nitrogen priorities of the cyanosphere and host cyanobacteria. Within the CAZy framework, the abundance of polysaccharide lyases in the cyanosphere is notable, as this may provide a mechanism for cyanobacterial EPS degradation or consumption ( 65 ). The EPS could provide a source of carbon for the cyanosphere. Polysaccharide lyases were found in 86% of the cyanospheres, highlighting the importance of this gene family within cyanospheres across cyanobacteria habitats. The key cyanosphere genera, Sphingomonas and Brevundimonas, are known for containing polysaccharide lyases. Sphingomonas strains can synthesize gellan lyase ( 66 ) and alginate lyases ( 67 ). Sphingomonas with polysaccharide lyase genes was found in 7 of the cyanospheres and was present in all terrestrial environments (1 subaerial, 1 hydroterrestrial, 5 soil). Perhaps its ability to degrade EPS contributes to its prevalence in all three terrestrial habitats. Brevundimonas , part of the “core” cyanosphere, also contained several polysaccharide lyases within 28% of the cyanospheres with the greatest abundance in Kastovskya adunca, Mojavia pulchra, and Nodosilinea sp. A particular group of polysaccharide lyases – alginate lyases– can degrade alginate, a polysaccharide found in algae and cyanobacteria ( 68 , 69 ). The cyanosphere contained six types of alginate lyases (PL6 alginate lyase, PL5 alginate lyase, M-specific alginate lyase, PL15 oligo-alginate lyase, PL17 alginate lyase, PL14 alginate lyase). Interestingly, polysaccharide lyases were also found in 42% of the cyanobacteria, with the most genes in Nostoc indistinguendum, Nostoc desertorum, and Pleurocapsa ercegovicii. Nostoc , in particular, is known for containing and utilizing polysaccharide lyases. Plant symbiotic Nostoc strains associated with feathermoss contained pectate lyases ( 70 ) which can be involved in plant pathogenesis and allow for symbiont-rhizobia infection ( 71 , 72 ). The pectate lyases found in feathermoss-associated Nostoc may assist with the colonization by Nostoc of the feathermoss. In the cyanosphere, this process would need to be reversed with various heterotrophic bacteria colonizing Nostoc . Implications The application of terrestrial cyanobacteria in ecosystem restoration is an emerging and rapidly expanding field ( 73 – 75 ). These microorganisms play a vital role in dryland ecosystem restoration due to their remarkable ability to withstand environmental stressors and the various ecological functions they support ( 76 ). Cyanobacteria contribute to critical processes such as stabilizing soils, enhancing soil fertility, promoting nutrient cycling, increasing water retention and infiltration, and sequestering carbon ( 74 , 75 , 77 , 78 ). These functions are essential for restoring degraded environments, particularly in arid and semi-arid regions where soil erosion, nutrient depletion, and poor water retention are prevalent challenges. The success of restoration efforts involving cyanobacteria can be closely tied to the cyanosphere ( 17 ). The composition of the cyanosphere may play a crucial role in determining the effectiveness of cyanobacteria in restoration activities. Therefore, it is essential 1) to investigate the relationships between cyanosphere microbes and their host more deeply, and 2) to consider the unique characteristics and requirements of the cyanosphere when selecting and applying cyanobacteria in ecosystem restoration projects to optimize their performance and ensure long-term ecological benefits. Data Availability The sequence reads for the project are deposited under BioProjects PRJNA1200267, PRJNA1208396, PRJNA1208397, and PRJNA1231202 (Table S1). Supplementary Table S1: Metadata for the cyanobacteria used in this study. The “Map ID” column corresponds with the locations of the cyanobacteria depicted in Figure 1 . The annual mean temperature and annual precipitation were derived from BioClim data when accurate latitude and longitude coordinates were provided for the cyanobacterial hosts. The * denote cyanobacteria that were not previously published in Ward et al. 2021. Table S2: Summary statistics from each MAG including completeness, contamination, number of contigs, genome size, and taxonomic information. Acknowledgments We are grateful to the National Park Service and Bureau of Land Management for granting us permission to study the cyanobacterial flora within Joshua Tree National Park (permit JOTR-2006-SCI-0018), Mojave Desert National Preserve (MOJA-2008-SCI-0024 and MOJA-2009-SCI-0039), and Grand Staircase-Escalante National Monument (permit UT-06-032-12-P). National Science Foundation (NSF) funded J.R. Johansen’s sample collection of soils from the Atacama Desert, isolation, and sequencing of those strains (NSF grant numbers DEB-0842702 and DEB-841734, respectively), as well as the collection and biodiversity research of biological soil crusts from various North American desert locations (NSF grant number DEB-9870201). This work is supported by the USDA National Institute of Food and Agriculture and Hatch Appropriations under Project #PEN04949 to E.C and Hatch project CA-R-PPA-5062-H to J.E.S. Any opinions, findings, conclusions, or recommendations expressed in this material are those of the authors and do not necessarily reflect the views of the National Science Foundation. The California Desert Research Fund at The Community Foundation, Robert Lee Graduate Student Research Grant, and the Phycological Society Grants in Aid of Research fund awarded to Nicole Pietrasiak provided support for the sampling campaigns and subsequent cyanobacterial research associated with strains from Joshua Tree National Park and Mojave Desert National Preserve. J.E.S. is a CIFAR Fellow in the program Fungal Kingdom: Threats and Opportunities. John Carroll University supported J.R.J.’s cyanobacterial culture collection since 1996. We greatly acknowledge the numerous students and colleagues who over the past 30 years collaborated with Johansen and led to the isolation of many of the strains investigated in this study. Funder Information Declared National Science Foundation , DEB-0842702 , DEB-841734 , DEB-9870201 Bureau of Land Management, https://ror.org/01sy5zn44 National Park Service, https://ror.org/044zqqy65 John Carroll University, https://ror.org/001gmya32 National Institute of Food and Agriculture , CA-R-PPA-5062-H , PEN04949 Phycological Society of America, https://ror.org/00a4fk439 References 1. ↵ Büdel B . 2011 . Cyanobacteria: Habitats and Species , p. 11 – 21 . In Lüttge , U , Beck , E , Bartels , D (eds.), Plant Desiccation Tolerance . 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