Bacterial Communities Associated with Poa annua Roots in Central European (Poland) and Antarctic Settings (King George Island)

Poa annua (annual bluegrass) is one of the most ubiquitous grass species in the world. In isolated regions of maritime Antarctica, it has become an invasive organism threatening native tundra communities. In this study, we have explored and compared the rhizosphere and root-endosphere dwelling microbial community of P. annua specimens of maritime Antarctic and Central European origin in terms of bacterial phylogenetic diversity and microbial metabolic activity with a geochemical soil background. Our results show that the rhizospheric bacterial community was unique for each sampling site, yet the endosphere communities were similar to each other. However, key plant-associated bacterial taxa such as the Rhizobiaceae family were poorly represented in Antarctic samples, probably due to high salinity and heavy metal concentrations in the soil. Metabolic activity in the Antarctic material was considerably lower than in Central European samples. Antarctic root endosphere showed unusually high numbers of certain opportunistic bacterial groups, which proliferated due to low competition conditions. Thirteen bacterial families were recognized in this study to form a core microbiome of the P. annua root endosphere. The most numerous were the Flavobacteriaceae, suspected to be major contributors to the ecological success of annual bluegrass, especially in harsh, Antarctic conditions.


Introduction
Poa annua L. (annual bluegrass) is one of the most ubiquitous grass species in the world. It can be found growing on every continent, having established populations even in the high Arctic and Antarctica [1][2][3]. Its extensive adaptability to a wide range of environmental conditions makes it a stubborn weed and pioneer species. P. annua has developed a series of features that ensure its ecological success. It can deal with biotic and abiotic stressors as well as habitat instability by compact growth habit, a high fraction of biomass allocated into belowground organs, and the presence of both annual and perennial life forms [4][5][6]. It also displays huge phenotypic and genotypic variability, making it difficult in the past to determine its evolutionary origin [1].
Base station, soil, mechanically altered by human activities; Skeletic Eutric Fluvisol (Turbic) The site is medium influenced by marine aerosols, moist, with large human influence, especially by using big vehicles.

Bacterial Extraction
Bacterial cells were extracted from fifteen (three per site) individual P. annua specimen rhizospheric soil and roots. The following method was devised based on the findings of [20] regarding the separation of prokaryotic cells from mineral and organic debris and the guidelines provided by [21] regarding root-associated microbe isolation. To analyze the microbiome of the root-adjacent soil, a sample of the soil was carefully removed from between the roots with a sterile spatula onto a pre-sterilized aluminum foil piece. Approx. 1 g of the soil was weighed and placed in a 50 mL conical tube containing 20 mL of sterile and cool (4 • C) dilution liquid composed of 0.9% (w/v) saline (NaCl) and 10 mM tetrasodium pyrophosphate (Na 4 P 2 O 7 ). The suspension was then shaken for 30 min in a TornadoTM Vortexer at 2000 rpm at 4 • C. The tubes were then placed in a VWR Ultrasonic Cleaner USC-TH filled with chilled water and sonicated for 60 s. The tubes were vortexed afterward for 30 s to suspend detached cells. After brief centrifugation (1 min; 1000 rpm; 4 • C), the suspension was submitted to metabolic fingerprinting by the Biolog Ecoplate technique. To detach the rest of the adhering soil, the root system was washed in 60 mL of sterile NaCl/Na 4 P 2 O 7 solution by shaking for 30 min in the aforementioned shaker (1000 rpm; 4 • C) and then rinsed 3 times in 5 mL of the same sterile and cooled solution by vortexing. Washed roots were sterilized by incubation in a cooled 10% hydrogen peroxide (H 2 O 2 ) solution for 5 min, then rinsed 3 times with sterile NaCl/Na 4 P 2 O 7 solution. The so surface-sterilized roots were placed in a pre-cooled sterile mortar. Two and a half ml of sterile NaCl/Na 4 P 2 O 7 was added with 0.6 g of sterile, sharp garnet sand (lysing matrix A) and gently ground with a pestle, allowing the sharp angular garnet pieces to comminute the roots to an amorphous Microorganisms 2021, 9, 811 4 of 19 pulp. The pulp was transferred to a 50 mL conical tube containing 20 mL of sterile and cool (4 • C) NaCl/Na 4 P 2 O 7 solution and submitted to the above-mentioned procedure (shaking, ultrasonication and vortexing). The resulting supernatant suspension was submitted to metabolic fingerprinting by the Biolog Ecoplate technique and DNA extraction.

DNA Extraction and Targeted 16S rRNA Gene Amplicon Sequencing
Rhizosphere soil DNA was extracted using the PowerSoil ® DNA isolation kit (QIA-GEN GmbH, Hilden, Germany) according to manufacturer protocol. An approx. 0.2 g of soil was used in triplicates. DNA solutions were kept at 4 • C for further analysis. The dilution liquid containing endosphere bacteria was passed through a sterile 47 mm Whatman polycarbonate filter (0.22 µm pore size). The DNA from the filter-trapped bacteria was extracted using the PowerWater ® DNA isolation kit (QIAGEN, GmbH, Hilden, Germany) according to manufacturer protocol and kept at 4 • C. This resulted in 72 DNA samples. The phylogenetic study was performed by targeted sequencing and analysis of the prokaryotic 16S ribosomal RNA gene. A fragment of the 16S rRNA gene containing the V3 and V4 variable regions was amplified using gene-specific primers: 16S_V3-F and 16S_V4-R positions 341-357F and 785-805R, respectively, according to Escherichia coli 16S rRNA gene reference sequence [22]. Illumina Nextera XT overhang adapter nucleotide sequences were included in addition to the 16S rRNA gene-specific sequences, which allowed sample indexing and pooling. Each PCR amplification was conducted in triplicates using KAPA HiFi PCR kit (Roche, Basel, Switzerland) in a final volume of 20 µL per reaction according to the manufacturer's instructions. Obtained PCR products were pooled into 10 samples (2 rhizocompartments × 5 sampling sites) in equimolar ratio and indexed using Nextera XT barcodes (Illumina, San Diego, CA, USA). Amplicon libraries were sequenced on Illumina MiSeq instrument (Illumina, San Diego, CA, USA) in the DNA Sequencing and Oligonucleotide Synthesis Laboratory (IBB, PAS). Sequencing was conducted in paired-end mode (2 × 300 bp) with the use of a v.3 (600 cycles) chemistry cartridge, which allowed the generation of long paired reads fully covering 16S V3-V4 amplicons.

Phenotype Fingerprinting with Biolog EcoPlate™
The EcoPlate Biolog assays assess the ability of a mixed microbial community to use any of 31 carbon compounds as the sole carbon source (plus a single control well with no-carbon). Microbial communities were characterized for their ability to catabolize 10 different carbohydrates, 9 carboxylic and acetic acids, 4 polymers, 6 amino acids, and 2 amines [23]. Root-associated bacterial suspensions were adjusted with sterile 0.9% saline to optical transmittance of 0.9. One hundred microliter aliquots of each suspension were added to each well of EcoPlate microplates (Biolog Inc., Hayward, CA, USA). The plates were incubated in darkness at 10 • C for Antarctic samples and 18 • C for European material. The temperatures were chosen to accommodate the activity range of the respective microbial communities: psychrophiles and psychrotrophes for Antarctic material, psychrotrophes and mesophiles for European material [24]. The color development (absorbance) was read at 590 nm (A 590 ) in a Varioscan plate reader (Thermofisher Scientific, Waltham, MA, USA), and cellular respiration was measured kinetically by determining the colorimetric reduction of tetrazolium dye. Data were collected approximately twice a week over a 65 day period. The prolonged incubation of EcoPlates was based on our previous observations [25][26][27]. Data from the forty-second day (Antarctic samples) and twenty-first day (European samples) of incubation were used as there was no further color development after this date. Final absorbance data were first blanked against the time zero reading and then blanked against the respective control well containing no-carbon source. Readings that had the A 590 value of 0.25 or higher were scored as a positive EcoPlate response (PER).

Measurement of Soil Components
Soil pH (in 1 M KCl) and salinity (in double-distilled water (ddH2O)) were measured with a CPC-411 Elmetron™ multiparameter probe according to [28]. Phosphates and ni-trates were determined spectrophotometrically in a Shimadzu UV 1601 spectrophotometer and in an Epoll-Eco 20 spectrophotometer respectively. Other elements were determined by atomic absorption spectroscopy [29].

Data Analysis
Raw sequencing data were cleaned, aligned, and classified automatically by the EzBio-Cloud platform using the PKSSU4.0 database [30]. Chimeric, low quality, and non-target (chloroplast, mitochondrial, and archaeal) amplicons were automatically discarded. The operational taxonomic unit was defined as a group of sequences that exhibit greater than 97% similarity to each other. Illumina reads were deposited in the NCBI Sequence Read Archive (SRA) as BioProject PRJNA678861. All results were compiled using Excel (MS Office) 2016 for Windows. A two-sample t-test was applied to compare different data sets. Variance within the sets was assessed using the f-test beforehand. Correlations between biological and geochemical parameters were calculated using Pearson's correlation coefficient. Principal component analysis was performed using the singular value decomposition method. Data visualization and statistical analysis has been performed using the R software (R v.4.0.2) and the following packages: ggplot2, fmsb, Hmisc, ggpubr, corrplot, and autoplot [31].

Diversity Indices
The highest diversity as assessed by operational taxonomic unit numbers (OTUs) were noted for the fertile Central European soil sample P2S (OTU = 7879), followed by Central European soil sample P1S (OTU = 6813). Antarctic soil samples displayed lower bacterial phylogenetic diversity. Sample P3S showed the highest diversity among them (OTU = 4498), whereas the proglacial soil sample P4S the lowest diversity (OTU = 1322). Bacterial communities residing within the roots displayed lower diversity (av. 2512.4 OTU, sd. 841) than the corresponding rhizosphere soil (av. 5165 OTU, sd. 2083). The most diverse was the bacterial community in soil sample P1R from grass specimens growing in nutrient-poor European soil (OTU = 3694). Samples P2R showed lower values (OTU = 2620). Root-residing bacteriome was diverse in plants growing near the Arctowski

Diversity Indices
The highest diversity as assessed by operational taxonomic unit numbers (OTUs) were noted for the fertile Central European soil sample P2S (OTU = 7879), followed by Central European soil sample P1S (OTU = 6813). Antarctic soil samples displayed lower bacterial phylogenetic diversity. Sample P3S showed the highest diversity among them (OTU = 4498), whereas the proglacial soil sample P4S the lowest diversity (OTU = 1322). Bacterial communities residing within the roots displayed lower diversity (av. 2512.4 OTU, sd. 841) than the corresponding rhizosphere soil (av. 5165 OTU, sd. 2083). The most diverse was the bacterial community in soil sample P1R from grass specimens growing in nutrient-poor European soil (OTU = 3694). Samples P2R showed lower values (OTU = 2620). Root-residing bacteriome was diverse in plants growing near the Arctowski Ant-

Bacterial Phylogenetic Diversity
Twelve major (>1%) phyla were found in the rhizosphere and root interior of the P. annua specimens ( Figure 2). Proteobacterial sequences were the most numerous in investigated samples. According to their percentage contribution, Proteobacteria have been enriched in the root-interior compartment in comparison to the respective rhizosphere soil community. The enrichment is approx. 2-fold in every case analyzed, being it European

Community-Level Physiological Profiling by Biolog EcoPlates
Microbial community response intensity in Biolog Ecoplates was considerably higher for the European samples than in those obtained from Antarctic material (Figure 4, Supplementary File S3). In order to be comparable, the data needed to be normalized in terms of intensity value (values within a particular sample type were divided by the highest value in that sample). The highest absorbance value at 590 nm (A590) obtained in the European material was A590 = 4.69, whereas, for the Antarctic samples, A590 = 2.82. D-mannitol was the most intensely metabolized carbon source by all examined microbial communities, especially by root-inhabiting communities from European material (A590

Community-Level Physiological Profiling by Biolog EcoPlates
Microbial community response intensity in Biolog Ecoplates was considerably higher for the European samples than in those obtained from Antarctic material (Figure 4, Supplementary File S3). In order to be comparable, the data needed to be normalized in terms of intensity value (values within a particular sample type were divided by the highest value in that sample). The highest absorbance value at 590 nm (A 590 ) obtained in the European material was A 590 = 4.69, whereas, for the Antarctic samples, A 590 = 2.82. D-mannitol was the most intensely metabolized carbon source by all examined micro-

Correlations between Biological and Geochemical Data
Pearson's correlation coefficient between the percentile abundance of bacterial families within the rhizosphere and the chemical composition of the soil revealed several significant (p < 0.05) negative correlations ( Figure 5A

Correlations between Biological and Geochemical Data
Pearson's correlation coefficient between the percentile abundance of bacterial families within the rhizosphere and the chemical composition of the soil revealed several significant (p < 0.05) negative correlations ( Figure

Principal Component Analysis
Principal component analysis (PCA) based on the abundance of sequences of a family-rank taxon showed a tight clustering of the root-dwelling bacterial communities, both of European and Antarctic origin ( Figure 6A). Soil bacterial communities were vastly different, with the proglacial sample P4S bearing the closest resemblance to root communities. PCA based on the Biolog Ecoplate responses showed that European root-interior microbial communities were the most similar despite the differences in their corresponding rhizospheric communities. Antarctic root communities bore more resemblance to their respective rhizosphere communities than to each other or the corresponding European samples ( Figure 6B). PCA of soil chemistry data showed a great discrepancy between the nutrient-poor (P1S) and the nutrient-rich European rhizospheric soils (P2S). Antarctic samples clustered closely together, showing great similarity ( Figure 6C). PCA clustering made using the combined phylogenetic and functional (Biolog Ecoplate) data revealed four distinct groups: European rhizospheric soil community (P1S, P2S), Antarctic rhizospheric soil community (P3S, P4S, P5S), European root community (P1R, P2R), and Antarctic root community (P3R, P4R, P5R) ( Figure 6D).

Principal Component Analysis
Principal component analysis (PCA) based on the abundance of sequences of a familyrank taxon showed a tight clustering of the root-dwelling bacterial communities, both of European and Antarctic origin ( Figure 6A). Soil bacterial communities were vastly different, with the proglacial sample P4S bearing the closest resemblance to root communities. PCA based on the Biolog Ecoplate responses showed that European root-interior microbial communities were the most similar despite the differences in their corresponding rhizospheric communities. Antarctic root communities bore more resemblance to their respective rhizosphere communities than to each other or the corresponding European samples ( Figure 6B). PCA of soil chemistry data showed a great discrepancy between the nutrient-poor (P1S) and the nutrient-rich European rhizospheric soils (P2S). Antarctic samples clustered closely together, showing great similarity ( Figure 6C). PCA clustering made using the combined phylogenetic and functional (Biolog Ecoplate) data revealed four distinct groups: European rhizospheric soil community (P1S, P2S), Antarctic rhizospheric soil community (P3S, P4S, P5S), European root community (P1R, P2R), and Antarctic root community (P3R, P4R, P5R) ( Figure 6D). Microorganisms 2021, 9, x FOR PEER REVIEW 11 of 19

Significant Differences between Microbial Parameters
The t-test calculations showcased significant (p < 0.05) differences between Central European and Antarctic rhizospheric microbial parameters (Figure 7). The relative abundance of the following taxa was higher in the European samples: Planctomycetes, Caulobacteraceae, Planctomycetaceae, Rhizobiaceae, Sinobacteraceae, Thermoleophilaceae, and Verrucomicrobiaceae. The catabolic intensity of several compounds was also higher in the Central European rhizosphere samples, most notably for: β-methyl-D-glucoside, phenylethylamine, glucose-1-phosphate, γ-hydroxybutyric acid, itaconic acid, D-galacturonic acid, and D-xylose. Antarctic material had a significantly higher sequence abundance of the Saccharibacteria phylum and the actinobacterial family of Microbacteriaceae. Significant differences between the root endosphere communities highlighted higher abundances/intensities of several features in the Central European samples. In those samples, a higher relative abundance of the following bacterial families were noted: Comamonadaceae, Cytophagaceae, Micromonosporaceae, Nocardioidaceae, and Rhizobiaceae. As for compound use intensity, it was true for the following substrates: β-methyl-D-glucoside, D-galactonic acid γlactone, D-galacturonic acid, L-asparagine, Tween 80, N-acetyl-D-glucosamine, γ-hydroxybutyric acid, D-cellobiose, glucose-1-phosphate, D-malic acid, and putrescine. The only feature significantly higher in the Antarctic material was i-erythritol use intensity. Most of the significant differences were apparent between the rhizosphere and the endosphere of P. annua. The root interior showed significant enrichment in the relative abundance of the following taxa: Proteobacteria, Microbacteriaceae, Sphingobacteriaceae, Hy-

Significant Differences between Microbial Parameters
The t-test calculations showcased significant (p < 0.05) differences between Central European and Antarctic rhizospheric microbial parameters (Figure 7). The relative abundance of the following taxa was higher in the European samples: Planctomycetes, Caulobacteraceae, Planctomycetaceae, Rhizobiaceae, Sinobacteraceae, Thermoleophilaceae, and Verrucomicrobiaceae. The catabolic intensity of several compounds was also higher in the Central European rhizosphere samples, most notably for: β-methyl-D-glucoside, phenylethylamine, glucose-1-phosphate, γ-hydroxybutyric acid, itaconic acid, D-galacturonic acid, and D-xylose. Antarctic material had a significantly higher sequence abundance of the Saccharibacteria phylum and the actinobacterial family of Microbacteriaceae. Significant differences between the root endosphere communities highlighted higher abundances/intensities of several features in the Central European samples. In those samples, a higher relative abundance of the following bacterial families were noted: Comamonadaceae, Cytophagaceae, Micromonosporaceae, Nocardioidaceae, and Rhizobiaceae. As for compound use intensity, it was true for the following substrates: β-methyl-D-glucoside, D-galactonic acid γ-lactone, D-galacturonic acid, L-asparagine, Tween 80, N-acetyl-D-glucosamine, γ-hydroxybutyric acid, D-cellobiose, glucose-1-phosphate, D-malic acid, and putrescine. The only feature significantly higher in the Antarctic material was i-erythritol use intensity. Most of the significant differences were apparent between the rhizosphere and the endosphere of P. annua. The root interior showed significant enrichment in the relative abundance of the following taxa: Proteobacteria, Microbacteriaceae, Sphingobacteriaceae, Hyphomicrobiaceae, Rhizobiaceae, Bacteroidetes, and Comamonadaceae while also displaying significantly lower catabolism intensities of L-phenylalanine, α-cyclodextrin, glycogen, and D-cellobiose. phomicrobiaceae, Rhizobiaceae, Bacteroidetes, and Comamonadaceae while also displaying significantly lower catabolism intensities of L-phenylalanine, α-cyclodextrin, glycogen, and D-cellobiose.

Poa annua Core Microbiome
Thirteen bacterial families had an average relative abundance higher than the average abundance of family-rank group in the root endosphere and are therefore considered to be the core microbiome taxa of P. annua in this study (

Poa annua Core Microbiome
Thirteen bacterial families had an average relative abundance higher than the average abundance of family-rank group in the root endosphere and are therefore considered to be the core microbiome taxa of P. annua in this study (

Discussion
The multiphasic approach applied in this study revealed several phenomena within the P. annua root-associated microbiome.
Rhizospheric soils examined in this study each contained a unique bacterial community in terms of phylogenetic diversity, as shown in the Principal Component Analysis. This diversity was higher in the European rhizospheric soil samples than in the Antarctic ones. Antarctic ecosystems were repeatedly proven to be less diverse and less complex than those in lower latitudes [32][33][34]. Geographical isolation was often made responsible for the low diversity of Antarctic macro-organisms [35,36]. However, this kind of isolation may not directly apply to bacteria, as it was discovered that live bacterial cells can be transported over long geographic distances [37]. Regarding rhizospheric microbial communities, their diversity can be largely controlled by host diversity and abundance [38], which in the case of Antarctic flora is indeed limited due to geographic isolation but also due to harsh climatic conditions [39]. The specific geochemical characteristics of the examined Antarctic soils may be even more responsible for the low bacterial diversity [40]. Those soils shared some features such as high salinity, sodium, magnesium, and heavy metal concentrations. This is mainly due to sea spray influence and the volcanic history of this site [41]. These characteristics, especially salinity, showed significant, negative correlations with members belonging to several soil community bacterial families, including the plant growth-promoting Rhizobiaceae. Salt heavy metal sensitivity, especially that of free rhizobial cells, has been extensively studied, as it impacts the biomass production of several agricultural plants [42][43][44]. Although there were no significant differences in the relative sequence abundance of the majority of bacterial phyla between European and Antarctic rhizospheric soils, they did significantly differ in the abundance of Planctomycetes and Saccharibacteria. European rhizosphere had a higher contribution of Planctomycetes, members of which are connected to plant biomass decomposition, but also to high soil calcium concentrations, which was noted in those soils [45]. Furthermore, European samples displayed significantly higher contributions of key taxa, such as the Rhizobiaceae, a family that contains plant-beneficial nitrogen fixers, and the Sinobacteraceae-harboring rhizospheric ammonia oxidizers [46,47]. The members of other families such as Caulobacteraceae and Verrucomicrobiaceae, known for grass-related rhizospheric compe-

Discussion
The multiphasic approach applied in this study revealed several phenomena within the P. annua root-associated microbiome.
Rhizospheric soils examined in this study each contained a unique bacterial community in terms of phylogenetic diversity, as shown in the Principal Component Analysis. This diversity was higher in the European rhizospheric soil samples than in the Antarctic ones. Antarctic ecosystems were repeatedly proven to be less diverse and less complex than those in lower latitudes [32][33][34]. Geographical isolation was often made responsible for the low diversity of Antarctic macro-organisms [35,36]. However, this kind of isolation may not directly apply to bacteria, as it was discovered that live bacterial cells can be transported over long geographic distances [37]. Regarding rhizospheric microbial communities, their diversity can be largely controlled by host diversity and abundance [38], which in the case of Antarctic flora is indeed limited due to geographic isolation but also due to harsh climatic conditions [39]. The specific geochemical characteristics of the examined Antarctic soils may be even more responsible for the low bacterial diversity [40]. Those soils shared some features such as high salinity, sodium, magnesium, and heavy metal concentrations. This is mainly due to sea spray influence and the volcanic history of this site [41]. These characteristics, especially salinity, showed significant, negative correlations with members belonging to several soil community bacterial families, including the plant growth-promoting Rhizobiaceae. Salt heavy metal sensitivity, especially that of free rhizobial cells, has been extensively studied, as it impacts the biomass production of several agricultural plants [42][43][44]. Although there were no significant differences in the relative sequence abundance of the majority of bacterial phyla between European and Antarctic rhizospheric soils, they did significantly differ in the abundance of Planctomycetes and Saccharibacteria. European rhizosphere had a higher contribution of Planctomycetes, members of which are connected to plant biomass decomposition, but also to high soil calcium concentrations, which was noted in those soils [45]. Furthermore, European samples displayed significantly higher contributions of key taxa, such as the Rhizobiaceae, a family that contains plant-beneficial nitrogen fixers, and the Sinobacteraceae-harboring rhizospheric ammonia oxidizers [46,47]. The members of other families such as Caulobacteraceae and Verrucomicrobiaceae, known for grass-related rhizospheric competence, have also been found in significantly higher amounts in those samples [48,49]. Antarctic rhizospheric soils had on average a higher contribution of Saccharibacteria sequences. Based on a limited amount of reports about the members of this phylum, they can be suspected to be scavengers or even parasites rather than active biomass degraders [50]. Members of the Microbacteriaceae displayed significantly higher amounts in the P. annua rhizosphere growing in the Antarctic. Rhizospheric enrichment in Microbacteriaceae has rarely been observed but could be influenced by climatic factors rather than soil composition [51]. A curious anomaly was noted in the Antarctic rhizospheric soils of postglacial origin. The postglacial rhizospheric soil sample contained high numbers of sequences belonging to the Firmicutes phylum, the majority of which were identified as Clostridiaceae, a family containing anaerobic, endospore-forming bacteria [52]. This can be connected to the recent deglaciation event [53]. Subglacial habitats have been speculated to be mostly anoxic. The experiment of [54] confirmed that in such conditions, members of the Clostridiaceae could be considerably enriched from Antarctic, supraglacial materials. Furthermore, the spores of the Clostridiaceae can persist in Antarctic settings even after the conditions have changed to oxic [26]. Besides higher phylogenetical diversity, European rhizospheric communities also displayed significantly higher use abilities for several types of compounds, such as carbohydrates, amino and carboxylic acids, lipids, and amines. Together with the high phylogenetic diversity, this suggests a plethora of different bacterial niches and high assemblage complexity, hinting at the presence of a rich reservoir of rhizosphere-competent bacteria [55,56].
The rhizosphere is considered the direct reservoir of root endophytes, albeit a selective entry mechanism exists and is well-described [57,58]. In this respect, the phylogenetic diversity in the endosphere was considerably lower than that in the rhizosphere. Furthermore, rhizosphere diversity had a direct impact on the endosphere diversity. This seems to be a staple when examining plant-associated bacterial communities as it was observed for wild plants [59] as well as a field [60] or greenhouse cultivars [61]. The endospheric communities of P. annua assessed in this study showed much phylogenetic similarity to each other when examined collectively with the rhizosphere community. This was probably due to the depletion in the root of several major soil-dwelling high-ranking taxa such as Acidobacteria and Planctomycetes, along with some other low abundance phyla. Sequences of the Proteobacteria and Bacteroidetes were significantly enriched in the endosphere. Despite this apparent similarity, several differences in the composition of the European and Antarctic endophytic communities could be noticed. Members of the family Rhizobiaceae and also Comamonadaceae were detected to largely contribute to European root endospheric bacteriomes but were significantly less numerous in Antarctic material. Those families hold nitrogen-fixing members, and their presence can largely increase plant biomass production [62,63]. In addition, the endosphere, as well as the rhizosphere of Antarctic P. annua specimens, were low in some of the most pivotal actinobacterial taxa found in well-developed soils in lower latitudes, namely the mycelium forming members of the families Nocardioidaceae and Micromonosporaceae, known for their antimicrobial activities [64,65]. On a family-rank level, there were no significantly enriched groups in the Antarctic endosphere. However, a few bacterial families showed abnormally high contribution in the Antarctic samples, namely: Oxalobacteraceae, Pseudomonadaceae, and Sphingomonadaceae. This is consistent with the findings of [66], where Pseudomonadaceae and Sphingomonadaceae were among high abundance taxa in the endosphere of native Antarctic plants: Colobanthus quitensis and Deschampsia antarctica. However, in those communities, the family Enterobacteriaceae was present in considerable amounts, unlike in the endosphere of examined here P. annua. The Pseudomonadaceae showed negative correlations with several key plant endophytic taxa such as the Rhizobiaceae, Intrasporangiaceae, Caulobacteraceae, and Sphingobacteriaceae, whereas the Oxalobacteraceae were negatively correlated with the Iamiaceae. Microbial interactions are usually very complex, albeit, within the root interior, they are more simplified due to the selection-mediated diversity impoverishment. In this respect, this could be a case of antagonisms or niche overlap. A recent study on the endophytic Oxalobacteraceae member Massilia sp. revealed its copiotrophic and r-strategy based lifestyle while also being competition-sensitive [67]. Such an explanation could be plausible in the Antarctic habitat, especially in the low diversity proglacial site, which houses opportunistic pioneer species [68]. The principal component analysis of the catabolic responses showed an interesting feature of the Antarctic bacterial endophytic community. While the Central European endophytic communities differed from the corresponding rhizospheric communities (and being more metabolically diverse), the Antarctic endophyte communities showed more similarity to the matching rhizospheric communities. This points toward Antarctic soil bacterial communities being catabolically fixed on certain abundant and stable nutrient sources, and the root-incorporated bacteria might not be true endophyte specialists. Furthermore, i-erythritol catabolism intensity was significantly higher in the Antarctic endophytic community. This sugar alcohol, contrary to the popular Mannitol, is not produced by plants but by green algae [69]. Antarctic soils house different species of aeroterrestrial algae, such as the widespread in the Antarctic region Prasiola crispa [10]. This compound's high catabolism potential in the Antarctic P. annua roots could mean aeroterrestrial algae-associated bacteria incorporation into the endophytic community. Despite the aforementioned dissimilarities between root communities, a core bacterial endophytobiome of P. annua can be extrapolated. The family Flavobacteriaceae has been on average the most abundant in the root interior, despite a wide abundance range. Only in recent years have the members of this family been recognized as important contributors to the root endosphere community [70] and were found to largely contribute to the endosphere of Antarctic native grass D. antarctica [71]. It is suspected that a metabolically diverse pool of Flavobacterium spp. is constantly present in the root interior, shifting in abundance patterns in response to environmental conditions and the physiological state of the host plant [70]. The latter is supported by the findings of [72], where plant-associated factors dictated flavobacterial presence. Investigations on temperate region invasive plants' root microbiota indicate that they increase the fitness of the host in non-native settings mainly by enhancing the acquisition of nutrients such as phosphorous and nitrogen [73]. However, in polar regions, the alleviation of cold-induced stresses may be of key importance for alien plant establishment [74]. In this respect, studies on the bacterially mediated cold resistance of agricultural crops point toward the action of ACC deaminase [75,76]. This microbially produced enzyme lowers the concentration of plant stress hormone ethylene, a substance that inflicts a significant reduction in plant growth and development [77]. Certain strains of the genus Flavobacterium have displayed ACC deaminase activity [78]. In consequence, Flavobacteriaceae bacteria may play a crucial role in P. annua adaptation to Antarctic conditions by modifying stress responses of the plant during adaptation [77].

Conclusions
In conclusion, bacterial root-associated communities of P. annua differed both phylogenetically and metabolically between those of Central European and those of maritime Antarctic origin. Several key plant-beneficial bacterial groups were less abundant in the Antarctic material, probably due to the geochemical makeup of the soil. Some bacterial families displayed unusually high abundance in the root endosphere of the Antarctic P. annua specimens. They most likely contained competition-sensitive opportunists that proliferated in the absence of antagonistic microbes. Nonetheless, an endophytic core microbiome could be assumed consisting of 13 bacterial families belonging to the Proteobacteria, Bacteroidetes, and Actinobacteria phyla. The Flavobacteriaceae family was the most numerous and most likely to positively influence the adaptation of P. annua to Antarctic conditions. Supplementary Materials: The following are available online at https://www.mdpi.com/article/10 .3390/microorganisms9040811/s1, Table S1: Alpha-diversity indices for the bacterial communities associated with Poa annua roots, Table S2: Relative abundance heatmap of sequences identified on a family-rank taxonomic level, Table S3: Heatmap displaying Poa annua rhizosphere and root community responses on Biolog Ecoplates, Table S4: Correlations of rhizospheric family-rank sequence abundance data and soil chemistry, Table S5: Correlations between root endosphere family-rank sequence abundance data.