Aerobic and anaerobic potential microbial consumption of salt marsh soils organic matter

doi:10.21203/rs.3.rs-2751051/v1

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Aerobic and anaerobic potential microbial consumption of salt marsh soils organic matter

https://doi.org/10.21203/rs.3.rs-2751051/v1

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Salt marshes have a high capacity to stabilize organic carbon in their soils as organic matter (OM). Globally, they are estimated to hold ~862 to 1,350 Tg of organic carbon in the upper meter. Which OM compounds are accumulated and which are the mechanisms protecting them, beyond anoxia, are important questions to unravel and understand the full potential of salt marsh ecosystems to store carbon. Soil microbial communities have a key role in OM degradation and, therefore, in its accumulation. In this study, performed at Ria de Aveiro (Portugal) coastal lagoon, two techniques were successfully applied to explore OM microbial mineralization in salt marsh soils: EcoPlates^TM under aerobic and anaerobic conditions, and Fourier-transform infrared spectroscopy (FTIR), together with principal component analysis on the transposed matrix, to extract the signal of specific compounds. The soil samples showed a high spatial homogeneity in microbial potential consumption, regarding both depth (up to 89 cm) and halophyte species. Potential aerobic consumption was high at all depths but the anaerobic communities showed a higher diversity in carbon sources consumption. Amino acids, carbohydrates and polymers were widely consumed, while half of the carboxylic acids were not consumed. Two of the carbon sources that were not consumed were likely protected within bigger molecules, D-galacturonic acids within hemicellulose and 4-hydroxy benzoic acids within phenolic compounds, being the last most likely accumulated as blue carbon. In addition, our study shows the potential of microbial incubations coupled with FTIR to greatly increase our understanding of salt marsh soils OM dynamics.

Blue Carbon

Biolog® EcoPlatesTM

Microbial activity

FTIR-ATR

Coastal wetlands

Coastal wetlands are among the most productive ecosystems on Earth, and bury and stabilize organic carbon as soil organic matter (SOM) [1, 2]. Vegetated coastal wetlands such as mangroves, salt marshes, and seagrass meadows, exceed the carbon sequestration rates of terrestrial forests, per unit area, by 1–2 orders of magnitude [2]. In salt marshes, > 98% of this organic carbon is stabilized in the soil [3]. Globally, they are estimated to hold ~ 862 to 1,350 Tg C in the upper soil meter [4]. SOM accumulation is promoted by their high primary production, capacity to bury allochthonous organic matter [1], and their prevailing anoxic conditions [5] due to daily tidal inundation. The composition and origin of the SOM influence its potential for accumulation [6] and mineralization upon environmental change [7]. Determining which mechanisms, beyond anoxia, promote the stabilization of SOM and what compounds are primarily stabilized is a necessary step to fully understand coastal wetlands SOM accumulation [8].

Soil microbial communities have a key role in SOM degradation and, therefore, in its accumulation [9]. Most studies focus on the capacity of microorganisms to promote CO₂ release through SOM mineralization, although considerable knowledge gaps still remain. Recent studies have also pointed out the role of microbial communities in producing stable SOM as a subproduct of their anabolism [10]. It has been hypothesized that those subproducts may be preferentially accumulated in aggregates or protected in mineral surfaces [11, 12]. Which types of plant and microbial molecules are accumulated as part of salt marsh SOM is one of the identified knowledge gaps in wetland microbial ecology and blue carbon science [8, 9].

Biolog® EcoPlate™ is an incubation method to characterize carbon source utilization by environmental microbial communities. Ecoplates contain thirty-one carbon sources (amines, miscellaneous compounds, polymers, carbohydrates, carboxylic acids and amino acids), a growth medium and a tetrazolium dye that develops color when the substrate is reduced. This method allows to examine which organic compounds are potentially consumed by microbial communities and, more relevant, which ones are not consumed and can potentially accumulate. Although Ecoplates are intended to measure aerobic degradation, they have been successfully utilized to measure anaerobic degradation potential of microbial communities in water [13] and of seagrass soil [14]. Attenuated total reflectance Fourier-transform infrared spectroscopy (FTIR-ATR), is a non-destructive analytical method used to infer the composition of a wide variety of materials. Recently, a new statistical protocol has been developed to measure the relative abundance of specific compounds in soil samples using their reference spectra [15, 16].

Here, the main aim was to assess the potential consumption of aerobic and anaerobic microbial communities along salt marsh soil cores, using Ecoplates, and to identify the presence of the Ecoplate carbon sources in the soil, using FTIR-ATR. The specific objectives of this study were: i) to assess which carbon sources are consumed by the soil microbial communities and which ones are not, ii) to assess differences in consumption between aerobic and anaerobic metabolisms; iii) to assess spatial differences (soil depth) and plant species-specificities in potential carbon consumption.

2.1. Sampling site and field methods

Ria de Aveiro (Portugal) is a mesotidal temperate coastal lagoon (45 km long; 10 km wide, Fig. 1) which intertidal areas range between 66 km² and 83 km² at low and high tide [17] and hosts mudflats, sandflats, seagrass meadows and extensive salt marshes [18, 19]. Its salt marshes are estimated to hold 216,346 Mg of organic carbon in the upper 25 cm of the soil: 69,017 Mg are stored under Juncus maritimus, 104,102 Mg under Atriplex portulacoides (formerly Halimione portulacoides), 16,524 Mg under Sarcocornia perennis and 6,461 Mg under Bolboschoenus maritimus [19].

Three vegetated, A. portulacoides (ATR), B. maritimus (BOL), and Juncus maritimus (J); and one bare sediment (BB) stations were selected. The stations were located close to each other to avoid differences in environmental conditions, like tidal inundation regime (Fig. 1). Three cores were retrieved per station (n = 3) using a Russian corer. The corer diameter was 4.5 cm and the length of the core was determined by the depth until the corer could be pushed manually: ATR 79 cm, BOL 89 cm, J 70 cm, and BB 50 cm max depth. The cores were sealed in PVC tubes with plastic film, transported to the laboratory and stored in darkness at 4 ºC until analysis.

2.2. Soil subsampling and laboratory analysis

The soil cores were sliced into 2 cm slices within an anaerobic chamber (Model 855-AC Plas Labs MI 48906 USA). In one core per station, samples for microbial incubation were selected equally spaced along depth: 2–4, 14–16, 24–26, 40–42, 52–54, 64–66, 74–76 and 84–86 cm. The upper (0 to 2 cm) and lower (deeper) part of each core were avoided as they were exposed to oxic conditions during retrieval. The same depths were subsampled in each core to allow comparisons among stations. Longer cores have larger number of samples. From the two replicate cores only two samples were retrieved for microbial incubations, at 2–4 and 40–42 cm depth.

Grain size distribution was determined at the same depths as microbial incubations by sequential sieving without SOM digestion, using a DIN ISO 3310-1 sieve set (Retsch AS 200 basic). Particle size was classified as: >2 mm, gravel; 2 − 1 mm, very coarse sand; 1-0.5 mm, coarse sand; 500 − 250 µm, medium sand; 250 − 125 µm, fine sand; 125 − 63 µm, very fine sand; and ≤ 63 µm, mud [20], and texture was classified following Blott and Pye (2012) [21]. From the longest core of each station, SOM and carbonate contents were analyzed every other sample and every 10 cm in the replicate cores. Organic matter and carbonate contents were estimated by Loss on Ignition technique (LOI%; 5 h combustion at 450°C and 2h combustion at 950°C, respectively).

Soil carbon sources’ composition was analyzed to assess the presence of the Ecoplates carbon sources in the soil, at the same depth as microbial incubations using a FTIR-ATR spectrometer (Agilent Cary 630) equipped with a single-reflection diamond crystal. Measurements were recorded in absorbance mode within the range of 4000 to 400 cm^− 1 at a spectral resolution of 2 cm^− 1, taking 100 scans per sample. A background was obtained before analyzing each sample. Reference spectra for the Ecoplates compounds were consulted at the SpectraBase library [22]. Only 20 compounds out of 31 were available: L-arginine, pyruvic acid methyl ester, D-xylose, D-galacturonic acid, L-asparagine, Tween 40, i-erythritol, 2-hydroxy benzoic acid, L-phenilalanine, Tween 80, D-mannitol, 4-hydroxy benzoic acid, L-serine, α-cyclodextrin, itaconic acid, glycyl-L-glutamic Acid, α-ketobutyric acid, phenylethyl-amine, D-mallic acid and putrescine.

2.3. Microbial incubations

All materials used were sterilized at 120°C for 1 h. Two samples (4 cm³ each), one for aerobic and one for anaerobic incubation, were taken from each section using a sterile polycarbonate cylinder, and stored in 50 ml falcon tubes with 30 ml of sterilized artificial seawater (formula: 25.9 g NaCl + 13.6 g MgCl₂ · 6H₂O + 4.14 g NaSO₄ in 1000 ml of H₂O MiliQ). The tubes were shaken at room temperature overnight to suspend and activate the microorganisms after the cold storage period. Suspensions were filtered through 15 to 20 µm filters (G43 Whatman).

Each Ecoplate well was inoculated with 100 µl of suspension using a multichannel micropipette and read right away to get the blank (T0, 590 nm, Biolog MicroStation™ Elx808BLG). The plates were then incubated at 25°C for 10 days and read every two days (T1-T4). Filtration and inoculation of anaerobic samples were performed in nitrogen atmosphere (Anaerobic chamber model 855-AC Plas Labs MI 48906 USA). The artificial seawater was bubbled with nitrogen (N₂) and the anaerobic tubes and inoculated plates were sealed with parafilm to avoid O₂ diffusion (see [14]).

2.4. Mathematical procedures

Data processing, statistical analyses and graphical representation were performed in R (v. 4.2.1) [23–30] and RStudio (v. 2022.11.0) [31]. Organic matter content data were log before testing for differences (p < 0.05; ANOVA and a post-hoc Tukey test). Carbonate content did not show normal distribution after transformation and therefore the non-parametric Wilcoxon tests was used instead.

The FTIR-ATR spectra were analyzed by Principal Component Analysis (PCA) using the transposed data matrix (i.e., wave numbers in rows, samples in columns), correlation mode and varimax rotation. With this approach, each sample spectrum is decomposed into scores' spectra each accounting for a given proportion of the sample's spectral variation. Five replicates of the available reference spectra of the compounds were also added as samples (in columns). Replication of the same spectra aims to reinforce its signal, forcing the PCA to extract it as a component (PC). The scores of the PCs obtained were cross checked with the reference spectra. Although the loadings of the PCs do not equate to the concentration of the compounds in the soil, they allow to check for relative changes among samples. This approach has been used before to study the composition of the organic and inorganic matrix of peat soils [15, 16].

The T0 absorbance values of the Ecoplates were subtracted to the absorbances after 10 days (T4). The average and coefficient of variation (CV) of the three replicates of each plate were calculated. Carbon sources with CV > 30 were examined to identify outliers. Outliers were not included in the final averaged absorbance of each carbon source per plate. For each plate, the average well color development (AWCD) was estimated as:

AWCD = Σ(R_i − C)/N (1)

where R_i is the color of each well at time i (after T0 subtraction), C is the color development in the blank (A1) and N is the number of carbon sources in the plate (31). The Shannon-Wiener diversity index (H’) was also calculated as:

H’ = −Σp_i(lnp_i) (2)

where p_i is the average corrected color of each well i divided by the sum of all the wells in the plate. Significant differences between metabolisms in AWCD and H’ were tested by t-test after testing for normality and variance homogeneity.

A global PCA with varimax rotation was performed with SOM and carbonates content and grain size fractions, aerobic and anaerobic AWCD and H’, and the relative abundance of those Ecoplate’s compounds detected in the soil, as per the transposed PCA loadings.

3.1. Geochemical parameters

Organic matter content ranged between 3.9 and 68.4% (Fig. 2A), being significantly lower at station BB (9.4 ± 5.8%, mean ± SD) and higher at station J (23.9 ± 11.3%, Fig. 2B). Carbonate content showed lower variability among stations, ranging between 2.2 and 21.9%, but followed a similar distribution, being significantly lower at station BB (4.8 ± 1.5%) and higher at vegetated stations: BOL 6.8 ± 3.3% (Fig. 2A). Both SOM and carbonate contents of the bare sediment were significantly lower than those of the vegetated stations (Fig. 2B). Samples showed textures from very slightly gravelly and slightly muddy sands to muddy sands; being very fine sand (125 − 63 mm) the predominant grain size (47.8 ± 8.2%, supplementary material Fig. S1).

The FTIR-ATR absorbance spectra and the second derivative spectrum showed the strongest absorbance and variation (measured as standard deviation) for bands in the region 1500 − 400 cm^− 1. The transposed matrix PCA yielded 17 PCs: 16 related to the reference spectra from the Ecoplates compounds (Table 1) and the first principal component likely related to the dominant inorganic matrix of the samples (Fig. 3; supplementary material Table 1, Fig. S2 to S17). Five of the 20 compounds included in the PCA showed negative loadings in the samples indicating that they were not detected in the soil (supplementary material Table S1 and Fig. S4, S5, S7, S10 and S17): α-Ketobutyric Acid, L-Phenilalanine, Itaconic Acid, 2-Hydroxy Benzoic Acid, and Glycyl-L-Glutamic Acid.

Table 1

Ecoplates compounds assigned to each principal component from the Fourier-transform infrared spectroscopy transposed matrix.
PCs	Main Compound	Secondary Compound
PC1	Inorganic matrix
PC2	i-Erithritol (C2)	D-Mannitol (D2)
PC3	Tween 40 (C1)	Tween 80 (D1)
PC4	Ketobutirc Acid (G3)	Pyrubic Acid Methyl Ester (B1)
PC5	L-Phenilalanine(C4)	Glycyl-L-Glutamic Acid (F4)
PC6	L-Asparagine (B4)
PC7	Itaconic Acid(F3)
PC8	Phenylethyl-amine (G4)
PC9	Putrescine (H4)
PC10	2-Hydroxy Benzoic Acid (C3)
PC11	D-Galacturonic Acid (B3)
PC12	D-Malic Acid (H3)
PC13	4-Hydroxy Benzoic Acid (D3)
PC14	D-Xylose (B2)
PC15	L-Serine (D4)
PC16	L-Arginine (A4)
PC17	Glycyl-L-Glutamic Acid (F4)

3.2. Microbial incubations

All amino acids, carbohydrates and polymers were consumed at all stations and redox condition (Fig. 4). One amine, one miscellaneous and five carboxylic acids showed low or not consumption at any depth or station. Carbon sources with similar color development to the control well were considered as not consumed, as color development in the control can be attributed to respiration of carbon sources present in the inoculum. There were significant differences between aerobic and anaerobic metabolism in both AWCD and H’. Aerobic metabolisms had a higher AWCD and lower H’ than anaerobic metabolism (Fig. 5). There were no significant differences among stations.

3.3. Spatial differences

The global PCA yielded 6 principal components explaining 85% of the total variance (Fig. 6; Table 2). The first principal component (PC1, 26%) clustered D-Galacturonic Acid, OM, Tween 40 and Tween 80, and i-Erithritol and D-Mannitol; against D-Malic Acid, Phenylethyl-amine, carbonates, Putrescine and the signal of the inorganic matrix. The second principal component (PC2; 22%) clustered 4-Hydroxy Benzoic Acid, L-Arginine, D-Xylose, L-Asparagine, and L-Serine against the signal of the inorganic matrix. The third principal component (PC3; 11%) clustered medium and fine sands against coarse sand. The fourth principal component (PC4; 11%) clustered Aerobic AWCD and aerobic and anaerobic diversity against very fine sands. The fifth principal component (PC5; 7%) clustered gravel and very coarse sand. The last principal component (PC6; 7%) clustered mud against anaerobic AWCD.

4.1. Carbon sources potential microbial consumption

Amino acids, carbohydrates and polymers were widely consumed in the studied soils. Those compounds were also highly consumed in other organic rich soils: under aerobic conditions in superficial (top 10 cm) samples of a bare sediment, A. portulacoides and S. maritima of the Ria de Aveiro salt marshes [32], a peat soil core [33], and soils under Posidonia australis and Posidonia sinuosa seagrass meadows [34]; and under both aerobic and anaerobic conditions in a Posidonia oceanica soil core [14]. A non-detectable potential consumption of a carbon source likely indicates that there isn’t a microbial community specialized in metabolizing that compound. Following the common approach in microbial biogeography that “everything is everywhere” but “the environment selects” [35, 36], the lack of a specialized community will indicate that the carbon source is not available in the soil or, in a competitive environment, microorganisms would have adapted to its exploitation. Four out of the seven carbon sources that showed no consumption were not detected in the soil samples, and for other two reference spectra were not available. Two compounds that did not show consumption at the Ecoplates were detected in the soil samples: D-galacturonic acid and 4-hydroxy benzoic acid, which could indicate that they are blocked for microbial consumption.

D-Galacturonic acid is usually present in soils as a component of hemicellulose [37]. In the current study it is more abundant in vegetated soils than in bare sediments, which agrees with a plant origin. It shows a steady decrease with depth in bare sediments and not clear patterns at the vegetated stations, likely due to the presence of roots (Fig. 3J). Hemicellulose is a complex molecule that usually requires a complex mix of enzymes for its degradation [38]. The decrease with depth at station BB (bare sediment) could indicate that hemicellulose is being degraded in the soil. However, we did not find a community specialized in D-galacturonic acid degradation in the incubations. This could indicate that, either the microbes able to degrade it cannot grow in laboratory cultures, or that D-galacturonic acid is not accessible in the soil, because the specific process of hemicellulose decomposition does not produce D-galacturonic acid as a subproduct [38]. Another far more common monomer of hemicellulose, xylose [38], showed very high consumption rates under both aerobic and anaerobic conditions (average 1.64 and 1.77 AWCD respectively), agreeing with a degradation of hemicellulose with depth.

A similar study on P. oceanica soils found that the 4-hydroxy benzoic acid was likely blocked from microbial access as part of a bigger recalcitrant plant derived phenolic compound [14]. Phenolic compounds are common in salt marsh soils as well [39, 40]. Furthermore, P-hydroxy benzoic acids can act as microbial inhibitors [41, 42], which may have prevented the consumption of this carbon source in our incubations.

It is interesting to notice that Tween 40 and 80 signals were detected in the soil (PC3, Fig. 3B). These compounds, are emulsifiers widely used in cosmetic and food industry. They are composed of derivatives of sorbitol (a sugar alcohol) and esterified fatty acids. Their presence in the soil could be related to contamination, but, more likely, the spectra detected could be derived from a composite signal of sugar alcohols and fatty acids. Forcing the PCA to look for the signal of specific compounds in our samples can lead to extraction of these composite signals, as long as the molecules combined are highly correlated. It is likely that both, fatty acids and sugar alcohols, are found in the labile SOM pool of the studied soils and show a similar distribution among soil depth and bare and vegetated stations.

4.2. Spatial differences

Aerobic and anaerobic consumption were observed at all depth and stations, signaling the presence of microorganisms with the capacity to metabolize SOM under both conditions. Ecoplates color development under aerobic consumption was observed in salt marsh, peat and Posidonia spp. meadows [32–34] and under aerobic and anaerobic conditions in a P. oceanica meadow [14]. All these environments are wetlands and, thus, water saturated at least for some time developing anoxic conditions. Nevertheless, there was potential aerobic consumption at all soils and depths, showing that if the soil was eroded and subject to oxic conditions, aerobic degradation of the accumulated SOM could begin without the need for a change in microbial community composition. This can be partially caused by salt marsh microbial communities functional redundancy, being many anaerobic microorganisms capable of aerobic respiration as well [9]. Nevertheless, salt marsh SOM decomposition after O₂ exposition is influenced not only by the microbial community, but also by SOM quality, showing faster decomposition rates at superficial, more fresh SOM rich, soil layers [7] despite the observed presence of a specialized aerobic community at all depths in this experiment.

In a P. oceanica soil core, where aerobic and anaerobic conditions were compared, aerobic consumption was observed at all depths but was predominant at the top 45 cm. Likely because of plant mediated O₂ diffusion to the rhizosphere [43]. Observable differences between rhizosphere and non-rhizosphere areas could only be visible if the O₂ release promotes a detectable increase in aerobic community size, which doesn’t seem to be the case in the current study. Salt marsh plant species are known to oxygenate their rhizosphere avoiding the formation of sulfides, but different aquatic plants had different O₂ transport efficiencies [44]. Furthermore, the cores studied here have a muddier texture compared to the core in the P. oceanica study [45], which may hinder O₂ diffusion, restricting microbial access to this electronic acceptor to a much smaller area around the rhizomes and roots, therefore, decreasing the size of the aerobic microbial community.

Despite the significant lower SOM content in bare sediment (station BB), there were no significant differences in AWCD nor in H’ index among stations after 10 days. Similar potential microbial consumption and diversity were found in bare and vegetated sediments. Our results disagree with those obtained in an aerobic Ecoplate incubation of superficial sediments of A. portulacoides, Spartina maritima and bare sediments, also from Ria de Aveiro salt marshes, in which bare sediments presented a lower AWCD than vegetated stations [32]. However, those incubations lasted for 36h while our first measurement was done at 62 h. There seems to be no differences between 62h (T1) and 10 days (230h, T4; Supplementary Material S18). If the first 30h showed differences in AWCD among stations, our experimental design could not register them.

Aerobic and anaerobic metabolisms were significantly different, with a higher AWCD but a lower diversity under aerobic conditions (Fig. 5A and 5B respectively). Aerobic microorganisms are more efficient than anaerobic ones in consuming SOM [46], explaining the higher AWCD in oxic conditions. The higher diversity under anoxic conditions could be related to the same process. As O₂ is a most energetic electron acceptor and aerobic microorganisms are more efficient consuming SOM, in a competitive environment with oxic and anoxic micro-niches, as salt marsh rhizosphere, the most energetic and easily consumed carbon sources will be likely utilized by aerobic organisms, while the anaerobic community may adapt to consume a wider range of carbon sources. Furthermore, oxic micro-niches in vegetated coastal wetland could be restricted to root tips [47], where root leaching of organic compounds provide fresh and easily degradable SOM, while anoxic conditions are predominant in the bulk sediment, consuming old buried SOM with a lower content of easily degradable compounds. This higher diversity in carbon sources consumption by the anaerobic community was observed in P. oceanica as well [14], suggesting similar microbial adaptations in blue carbon ecosystems.

Global PC1 shows a separation between bare and vegetated stations (Fig. 6A). On one hand, it clusters Tween 40, Tween 80, i-Erithritol, D-Mannitol with OM content. i-Erithritol and D-Mannitol are carbohydrates and the signal of Tween 40 and 80 was discussed above as being a composite signal of labile SOM compounds. On the other hand, it clusters Putrescine, Phenylethyl-amine, and D-Malic Acid with the inorganic matrix signal. Both Putrescine and Phenylethyl-amine are important molecules produced by plant and microorganisms, and Malic acid is a known rhizosphere exudate [48]. However, they show a stronger signal in unvegetated samples. Although they are common molecules, they are found in much lower concentrations than other organic compounds, such as cellulose and lignin. Therefore, in vegetated areas their signal may be diluted by other organic compounds.

The global PC2 separated ATR2-4, ATR64-66, J14-16, J24-26, and J52-54 from the other samples (Fig. 6A). Those samples contained large roots. Both 4-Hydroxy Benzoic Acid, as part of a phenolic compound, and D-Xylose are likely to be part of structural plant molecules like lignin and cellulose [38, 40]. Rhizosphere leachates usually have a high concentration of organic nitrogen-rich molecules [10], which may explain the correlation between L-Arginine, L-Asparagine and L-Serine and structural plant molecules. Therefore, PC2 could be an indicator of root presence. Both PC3 and PC5 indicate differences in soil texture with no clear differentiation between stations. PC4 and PC6 show negative relations between potential microbial activity and very fine sands and mud. Both PCs could be showing the role of fine fractions in protecting SOM, preventing microbial access. Color development in Ecoplates is influences by the initial size of the inoculum [49]. Thus, the SOM content being equal, samples with higher content of fine fractions will prevent microbial access to part of that SOM, limiting the size of the microbial community, and therefore Ecoplates color development.

4.3. Conclusions and final remarks

The soil cores selected for this study, bare sediment and soils vegetated by B. maritimus, A. portulacoides and J. maritimus, showed high homogeneity in SOM potential microbial consumption among the stations and with depth. Potential aerobic carbon consumption was high at all depths despite the anaerobic nature of these soils, showing the high potential for the mineralization of the accumulated SOM stocks if the soil is eroded and oxygenated. The anaerobic communities showed a higher diversity of potential carbon sources consumption, likely due to a lack of easy degradable carbon sources in the environment. Amino acids, carbohydrates and polymers were widely consumed, while half of the carboxylic acids were not consumed. Most of the carbon sources that were not consumed were absent in the soil, therefore the microbial communities were not adapted to their exploitation; while two other carbon sources were likely protected within bigger molecules, D-galacturonic acids within hemicellulose and 4-hydroxy benzoic acids within phenolic compounds, being the last quite likely accumulated in the soil.

Coupled analysis of SOM composition and microbial incubations is a promising approach to better understand microbial mineralization dynamics. In this study, two techniques were successfully applied to explore this process: Ecoplates under aerobic and anaerobic conditions and FTIR-ATR, together with PCA on the transposed matrix to extract the signal of specific compounds. As Ecoplates are a commercial kit, the carbon sources tested are not specific of salt marsh soils. Incubations for specific salt marsh organic compounds along environmental gradients will greatly increase our understanding of their SOM dynamics.

Acknowledgments

The authors would like to thank Amadeu Soares for providing access to essential equipment, Abel Ferreira for his assistance installing the nitrogen chamber and Mariana Caramujo for her support in field and laboratory analysis. The study was also supported by “Grupos de Referencia Competitiva” (ED431C 2021/32) by Xunta de Galicia and the project MARSH-C-LEVEL (CNRS/INEE, via OHMI – “Centre National de la Recherche Scientifique”, via International Observatory Hommes-Millieux). NP-J was supported by the Programme for Requalification of the Spanish University System 2021-2023, modality Margarita Salas. AIS was supported by national funds through the FCT-Foundation for Science and Technology, I.P., under the project/contract CEECIND/00962/2017. NP-J and AIS also thank FCT/MCTES for the financial support to CESAM (UIDB/50017/2020 + UIDP/50017/2020 + LA/P/0094/2020), through national funds.

Funding

The study was supported by “Grupos de Referencia Competitiva” (ED431C 2021/32) by Xunta de Galicia and the project MARSH-C-LEVEL (CNRS/INEE, via OHMI – “Centre National de la Recherche Scientifique”, via International Observatory Hommes-Millieux). NP-J was supported by the Programme for Requalification of the Spanish University System 2021-2023, modality Margarita Salas. AIS was supported by national funds through the FCT-Foundation for Science and Technology, I.P., under the project/contract CEECIND/00962/2017. NP-J and AIS also thank FCT/MCTES for the financial support to CESAM (UIDB/50017/2020 + UIDP/50017/2020 + LA/P/0094/2020), through national funds.

Conflicts of interest/Competing interests

The authors declare no competing interests.

Ethics approval

No ethical approval was required.

Consent to participate and consent for publication

No approval was required to accomplish the goals of this study because the experimental work was conducted with environmental microbial communities.

Availability of data, material and code availability

The data and code used in this manuscript can be consulted at the GitHub page of the corresponding author (https://github.com/NPJuncal/Ecoplates2022)

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Aerobic and anaerobic potential microbial consumption of salt marsh soils organic matter

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Abstract

Figures

1. Introduction

2. Material And Methods

2.1. Sampling site and field methods

2.2. Soil subsampling and laboratory analysis

2.3. Microbial incubations

2.4. Mathematical procedures

3. Results

3.1. Geochemical parameters

3.2. Microbial incubations

3.3. Spatial differences

4. Discussion

4.1. Carbon sources potential microbial consumption

4.2. Spatial differences

4.3. Conclusions and final remarks

Declarations

References

Additional Declarations

Supplementary Files

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