A Protocol for the Characterization of Diatom Communities in Mountain Glaciers

Abstract

This research presents a significant contribution to the methodologies and protocols for studying diatom communities in cryoconite holes on glaciers. Cryoconite holes are unique microenvironments found on glacial surfaces that support intricate microbial ecosystems, with diatoms playing a pivotal role in these communities. The refined methodologies developed in this study include optimizing sampling techniques to ensure that collections are both representative and diverse, which is crucial for accurate ecological assessments. Additionally, advanced digestion processes have been implemented to effectively isolate and purify diatom samples while minimizing contaminants, thereby improving sample integrity. Improved microscopic mounting techniques enhance visual clarity, facilitating more precise identifications of diatoms under the microscope. Furthermore, integrating DNA-based taxonomy broadens the taxonomic scope, providing valuable molecular insights into the diversity and evolutionary relationships of diatoms. Collectively, these protocols aim to enhance the reliability, depth, and multidimensional understanding of diatom ecology in cryoconite holes and broader glacial ecosystems, ultimately contributing to the field of glaciology and microbial ecology.

1. Introduction

Cryoconite holes are depressions found in the ablation zones of glaciers worldwide. These depressions are typically cylindrical and trap wind-blown sediments [1]. Often isolated from the atmosphere and the surrounding supraglacial hydrological system, they are influenced by various abiotic and biotic factors [2].

Cryoconite holes, ice-bound habitats on glaciers, harbor diverse microbial communities adapted to extreme conditions. These communities vary between Arctic and Antarctic regions, with distinct bacterial and microalgal compositions [3]. In Antarctic cryoconite holes, diatoms from genera such as Muelleria and Diadesmis are prevalent, with community composition influenced by location and environmental factors [2]. In contrast, Andean glaciers exhibit rare but diverse diatom assemblages, dominated by cosmopolitan and aerophilic species [4]. Interestingly, cryoconite holes with higher biodiversity and biomass show less partitioning between sediments and water columns, potentially due to mixing or freezing effects [5]. Geographic location, environmental conditions, and glacier characteristics all contribute to the unique microbial habitats of cryoconite holes, adding to the overall biodiversity of glacial ecosystems.

Studies reveal that cryoconite holes can harbor distinctive diatom communities influenced by factors such as location, freeze–thaw conditions, and sediment characteristics [2]. Diatom genus richness in cryoconite holes can be comparable to or higher than in adjacent aquatic habitats, suggesting multiple sources for colonization, including terrestrial environments [6,7]. The diatom flora in cryoconite holes shares similarities with Arctic lakes and moss communities, indicating potential for seeding various habitats as glaciers retreat [7]. Bacterial communities in these holes, dominated by Proteobacteria and Cyanobacteria, show strong correlations with metabolite profiles and exhibit significant differences between alpine and Arctic regions, reflecting local and regional influences on microbial ecosystems in glacial environments [8].

To characterize diatom communities in mountain glaciers, a suitable protocol involves leveraging high-throughput sequencing methods tailored to extreme environments like glacier-fed streams (GFSs) [9]. This protocol must consider the low biomass and high inorganic particle load in GFS sediment samples, which can affect nucleic acid extraction efficiency. An adapted phenol–chloroform-based extraction method has been shown to yield higher DNA quantities and accurately recover taxonomic profiles and genomes of microbial communities, including eukaryotes, in glacier-fed streams [9]. Additionally, diatom diversity in high-altitude glaciers, such as the Sach Pass glacier in India, highlights the need for selecting robust diatom communities capable of surviving harsh temperature conditions at higher altitudes [10]. By combining insights from these studies, a comprehensive protocol for diatom community characterization in mountain glaciers can be developed, ensuring efficient nucleic acid extraction and accurate representation of microbial diversity.

Recent studies have explored diatom communities in Ecuadorian mountain glaciers and rivers, revealing their potential as bioindicators of environmental change. Cryoconite holes on Antisana Glacier harbor diverse diatom assemblages, with elevation driving community structure [10]. In the Pita River, epilithic diatom communities show high species richness and respond to eutrophication gradients, though their trophic values may differ from those reported in the literature [11]. Genetic barcoding of Ecuadorian diatoms has identified species suitable as water quality bioindicators, including Sellaphora seminulum, Nitzschia fonticola, and Achnanthidium minutissimum [12]. These studies contribute to the development of diatom-based bioassessment tools for Ecuadorian aquatic ecosystems, highlighting the need for region-specific indices and the potential of combining morphological and molecular approaches for diatom identification and ecological assessment.

In the scientific context of Ecuador, research related to diatoms in glaciers is virtually non-existent. Our work stands as a pioneering initiative in this field, aiming to fill this knowledge gap in Ecuadorian glacial ecosystems. This study seeks to significantly contribute to the understanding of diatom ecology in these unique and extreme environments.

Cryoconite holes are known to host diverse microbial communities, including diatoms. A highly diversified diatom community in cryoconite holes through microscopic analysis has been documented [10], and Wang et al. [13] investigated microbial communities using amplicon sequencing of the 16S rRNA gene. Novotná Jaroměřská et al. [14] described spatial and temporal patterns in diatom biomass and community structure in cryoconite from the ablation zone of a glacier, and Vinšová et al. [6] found that diatom communities in these holes differ significantly from those in nearby lakes, suggesting a unique source of species. These papers provide valuable insights into the methods used to characterize diatom communities in cryoconite holes of glaciers, underscoring the need for comprehensive and dynamic methods to consider both unique sources and environmental influences.

To collect samples from cryoconite holes, researchers typically follow unsystematic approaches. Studies in the Central Italian Alps, Arctic, Norway, the Alps, and maritime Antarctic have employed methods involving the division of cryoconite holes into sampling zones based on water flow on glaciers, with subsequent collection of subsamples from different zones within the holes [14,15]. The collection process usually includes the extraction of cryoconite material from the ablation zone of glaciers over entire ablation seasons to capture spatial and temporal patterns in biomass, community structure of photoautotrophs, invertebrates, stable isotopic composition, and organic matter content [14]. Researchers usually analyze sediments and water columns separately to study spatial partitioning of microbial communities [5]. Various techniques are employed to characterize these ecosystems, including DNA extraction, chlorophyll a quantification, and geochemical analyses such as 13C and 14C measurements [16].

Given the absence of previous systematic studies, it is crucial to address this issue, especially in tropical glaciers like those in Ecuador, which present distinctive environments with unique conditions for microscopic life. We aim to present a simple and effective methodology that facilitates the collection and analysis of diatom data in these glaciers, serving as a reference for future research in this emerging field.

2. Materials and Methods

2.1. Preliminary Sampling of Diatoms in Glaciers

With the aim of determining the presence of diatoms in the glacier ablation zone in Ecuador, preliminary sampling was conducted, collecting samples from three specific locations: (a) white ice without sediment, (b) the surface of the cryoconite hole itself, containing sediment deposited by aeolian processes, (c) the cryoconite hole containing water and sediment (Figure 1a–c).

2.2. Study Area and Sample Collection

The tropical glaciers studied in Ecuador are associated with the Antisana, Cotopaxi, Cayambe, and Carihuairazo volcanoes, as illustrated in Figure 1d. These glaciers are located in varied geographic contexts, enabling an examination of their unique characteristics and the assessment of their relative significance. Initially, sampling was conducted on the Antisana Glacier using monthly transects of 200 m along the altitudinal gradient of the glacier (1 day/month), ascending to the summit or until no cryoconite holes were found. Samples were collected from all cryoconite holes along this transect, irrespective of their size. Subsequently, field campaigns lasting three to five consecutive days were conducted for the other glaciers and again in Antisana, with samples collected randomly from cryoconite holes from the summit downwards. The sampling duration on each glacier was determined based on weather conditions and the time required to cover the entire altitudinal gradient.

All cryoconite holes were in a frozen state. After breaking the ice layer, sediment and water were extracted using a 50 mL syringe and stored in sterilized 100 mL bottles with Transeau solution (consisting of deionized water, ethanol, and formalin in a 6:3:1 ratio). These samples were used for characterization through light microscopy and scanning electron microscopy (SEM) when necessary. Samples for metabarcoding were extracted using another 50 mL syringe and stored in sterilized 50 mL vials. The collected and labeled samples were placed in sterilized ziplock bags inside a cooler at −4 °C in complete darkness. Subsequently, these samples were transported to the Limnology Laboratory of the Faculty of Engineering and Applied Sciences of the SEK Private University, following the methodology previously described by [10]

In each cryoconite hole, measurements of length, width, and depth were taken, along with various physicochemical variables, including conductivity, pH, alkalinity, nitrites, nitrates, dissolved oxygen, and temperature. These measurements were carried out using a HANNA HI9829 multi-parameter(HANNA instruments, Woonsocket, RI, USA) and the “Easy Strips” field kit from the Tetra brand, following the procedures outlined in the study by [10].

2.3. Sample Pretreatment

The sediment samples collected from the site revealed a substantial presence of non-organic volcanic material, which demonstrated a remarkable resistance to conventional digestion methods [11,21,22,23,24]. To address this challenge, a filtration process was implemented using an entomological mesh with a porosity of 100 µm (Entomopraxis, Barcelona, España) which was previously sterilized with UV. This technique enabled the retention of coarse volcanic material while allowing the passage of diatoms. The procedure involved placing the mesh at the top of a precipitation beaker, adding the sediment, and using distilled water to facilitate the passage of the material. Upon completion of filtration, 50% of the treated sample intended for genetic analysis was extracted and stored in 50 mL Falcon tubes. This sample underwent centrifugation at 4000 rpm for 5 min, preserving the resulting sediment. The remaining 50% underwent digestion processes for subsequent optical analysis.

2.4. Molecular Analysis

2.4.1. DNA Extraction and Quantification

The samples for molecular analysis were processed using the DNeasy PowerSoil^® Pro Kit (Qiagen, Hilden, Germany) for the extraction process following the protocol used by [25], with certain modifications to the procedure focusing on supplementing the lysis stage. This involved adding 800 μL of CD1 solution provided by the kit to the PowerBead Pro Tubes (Qiage, Hilden, Germany) and leveling with sediment from the previously treated samples, followed by vortexing for 30 min and heating at 60 °C for 15 min, with intermittent vortexing for 5 s every 5 min.

The protocol was strictly followed from step 3 to 15 based on the manual from the commercial supplier [26], leading to the final collection stage where the quantity of C6 solution was replaced with 30 μL, incubating for 5 min at room temperature (15 to 20 °C), followed by centrifugation at 15,000× g for 1 min. The DNA was stored in 2 mL Eppendorf tubes along with the corresponding identification code. DNA quantification was performed using a Qubit™ dsDNA HS Assay (Thermo Fisher Scientific, MA, USA).

2.4.2. DNA Integrity Control

The integrity of the extracted DNA was verified using agarose gel electrophoresis with a 2% concentration in TAE buffer, employing SafeView™ Classic (Richmond, BC, Canada) as the staining agent. Five microliters of the sample were run alongside 2 μL of loading buffer under a steady current of 100 V for a duration of 60 min.

2.4.3. Library Preparation and Sequencing

For library construction, a 312 bp fragment of the rbcL gene was amplified using an equimolar mix of primers proposed by [27] (

Table 1. The rbcL primers used for amplification [27].

Primer	Sequence (5′ to 3′)
Forward—Diat_rbcL_708F_1	AGGTGAAGTAAAAGGTTCWTACTTAAA
Forward—Diat_rbcL_708F_2	AGGTGAAGTTAAAGGTTCWTAYTTAAA
Forward—Diat_rbcL_708F_3	AGGTGAAACTAAAGGTTCWTACTTAAA
Reverse—Diat_rbcL_R1	CCTTCTAATTTACCWACWACTG
Reverse—Diat_rbcL_R2	CCTTCTAATTTACCWACAACAG

), incorporating Illumina adapters at the 5′ end: P5 (CTTTCCCTACACGA CGCTCTTCCGATCT) and P7 (GGAGTTCAGACGTGTGCTCTTCCGATC).

In the first amplification step, PCRs were performed in a final volume of 12.5 μL, containing 1.25 μL of DNA, 0.5 μM primer mix, 3.13 μL of Supreme NZYTaq (Lisboa, Portugal) 2x Green Master Mix, and ultrapure water to 12.5 μL. The samples underwent thermocycling under the following conditions: denaturation at 95 °C for 5 min, 35 cycles of 95 °C for 30 s, 46.3 °C for 45 s, 72 °C for 45 s, and a final extension step at 72 °C for 7 min. Each sample was run in triplicate.

In the second amplification step, the PCR mix contained 2.5 µL of DNA, 0.5 µM of primers, 6.25 µL of Supreme NZYTaq 2x Green Master Mix (NZYTech), CES 1X [28], and ultrapure water to a final volume of 12.5 μL. Oligonucleotide indices required for multiplexing different libraries in the same sequencing pool were added in a second amplification step under identical conditions but with only 5 cycles and an annealing temperature of 60 °C. A negative control containing no DNA was included in each PCR cycle to verify contamination during library preparation.

Library sizes were confirmed using 2% agarose gels stained with GreenSafe (NZYTech, Lisboa, Portugal) and visualized under UV light. Subsequently, libraries were purified using Mag-Bind RXNPure Plus magnetic beads (Omega Bio-tek, Georgia, USA), following the manufacturer’s instructions. Final libraries were pooled in equimolar amounts based on quantification results obtained using the Qubit dsDNA HS double-stranded DNA assay (Thermo Fisher Scientific). The pool was sequenced in a fraction (3/16) of a MiSeq PE300 flow cell (Illumina, CA, USA).

2.4.4. Data Quality Control

The paired-end file corresponding to the Illumina sequencer libraries consists of forward (R1) and reverse (R2) reads, each of which includes its respective quality scores. Sequence adapter trimming was performed using the Cutadapt v3.5 tool [29]. The sequencing read quality was assessed with the FastQC program [30], and the results are summarized using MultiQC [31].

2.4.5. Bioinformatic Analysis

For the integration of sequencing reads, the QIIME 2 program (version 2023.2) [32], was employed using the DADA2 tool [33]. Firstly, primer sequences for PCR and reads with high interference and low quality were removed. Subsequently, data dereplication or combination of similar R1 and R2 reads was performed, merging sequences with a minimum of 12 aligned base pairs. Finally, a list of amplicon sequence variants (ASVs) was generated, and chimeric sequences were filtered out. As a result of the bioinformatic analysis, occurrences of each ASV in each sample are tabulated, along with their most representative sequences, which will be used for their respective taxonomic assignment.

2.4.6. Taxonomic Assignment

Taxonomic assignment was performed using the Diat.barcode database [34] [version 11.1, May 2022] as a reference. The representative sequences of each ASV are aligned using a classify-sklearn algorithm implemented in a QIIME 2 program. Each ASV is tabulated with its respective taxonomic information, and assignments are filtered based on the following parameters: singletons or sequences with frequencies less than 0.01% in each sample are excluded. Similarly, sequences not assigned to the phylum Bacillariophyta or unidentified sequences are discarded.

2.5. Microscopic Identification

2.5.1. Sample Digestion

The digestion process of Antisana Glacier samples, as described by [10], followed the protocols outlined in [11,21,22,35]. The work can be performed with limited materials or items, namely 30% hydrogen peroxide, a hot plate, optical microscope, and a centrifuge. To achieve frustules completely devoid of organic matter, the samples, which had been previously filtered and placed in test tubes, underwent a digestion process using only 30% hydrogen peroxide. These test tubes were arranged in a pot filled with sand and positioned on a hot plate, maintaining a constant temperature of 100 °C for an uninterrupted period of 48–92 h. Hydrogen peroxide is added as needed during this process.

Once the total absence of organic matter in the sample was confirmed, the sample was subjected to a decanting or centrifugation process to recover the resulting sediment. The sediment was then stored in labeled and sterilized glass vials.

2.5.2. Sample Concentration

Due to the low abundance of individuals in the glacier samples, a rigorous procedure was required to obtain a representative sample. Each sample vial was vigorously shaken and then subjected to a 10 s time count after homogenization. Subsequently, using a Pasteur pipette, 1 mL of the resuspended material was extracted, avoiding sediment removal. This process was repeated ten times to effectively concentrate the sample. The resulting concentrated sample was placed in Eppendorf tubes and finally examined under an optical microscope at 400× magnification to verify the presence of diatoms. Alternative methods for valve concentration in similar scenarios have been published [36,37,38,39].

2.5.3. Taxonomic Identification

The permanent microscopic slides were analyzed using a Euromex PC/, Duiven.

The Netherlands DIC optical microscope equipped with a CMEX-Pro-18 camera, allowing for observation at a magnification of 1000×. Taxonomic identification was based on the analysis of type material in specialized literature such as [40,41,42,43,44,45,46,47,48,49,50,51,52,53,54,55,56] and websites such as [57,58,59].

The methodological approach is based on the proposal by [23], adapting the protocol for samples with low abundance, which involves a comprehensive scan of the slide to record the highest number of individuals. Diatoms were captured in photographs using the digital extension Image Focus Alpha for Windows v1.3.7.22259 and processed using CorelDRAW Graphics Suite 2022 v24.3.0.571 Multilingual software, generating identification matrices. The process described above is illustrated in Figure 2.

3. Results and Discussion

Conducting the pre-sampling in the Ablation Zone of the glacier was crucial for accurately determining the specific locations of diatom presence. This analysis revealed that in mountain glaciers, diatoms are exclusively found in cryoconite holes. This result is consistent with findings from other studies, highlighting these ecosystems as habitats where this microalga is present [2,7,60,61,62].

The use of an entomological mesh with a porosity of 100 µm before the digestion process, as well as the concentration of samples after digestion, proved to be essential and highly efficient in this ecosystem. This approach successfully eliminated a significant portion of large and coarse volcanic material, thereby facilitating the mounting of slides and enhancing the observation of species during identification (Figure 3). The same sample treatment process before and after the digestion procedure applied to the Cotopaxi Glacier was replicated in the Cayambe and Carihuairazo glaciers, yielding identical results.

Comparison of Morphological and Barcoding Inventory on the Antisana Glacier

While this methodology was applied to all four glaciers—Antisana, Cotopaxi, Cayambe, and Carihuairazo—we will focus specifically on presenting the results from the Antisana Glacier.

A total of 57 taxa (160 individuals) were identified using light microscopy, including genera, species, subspecies, and varieties. The taxa with the highest relative abundances were Hantzschia amphioxys (Ehrenberg) (6.9%), Gomphonema parvulum (Kützing) (6.9%), Pinnularia borealis Ehrenberg (4.4%), Pinnularia sp2 (Ehrenberg) (4.4%), Pinnularia borealis var. islandica Krammer (3.8%), and Pinnularia borealis var. lanceolata Hustedt (3.8%).

In contrast, 10 taxa (86,287 individuals) were identified through Illumina sequencing, which allowed classification at the order, family, genus, and species levels. The taxa with the highest relative abundances using this method were Psammothidium germainii (Manguin) Sabbe (97.23%), Rhoicosphenia abbreviata (C.Agardh) Lange-Bertalot (1.87%), Naviculales sp. pl. (0.53%), and Bacillariophyceae sp. pl. (0.24%).

When analyzing the ecological parameters of the diatom communities identified by light microscopy (Shannon 3.758, Chio-1: 70.04) and metabarcoding (Shannon 0.155, Chio-1: 10), the results suggest that microscopy provides a more complete picture of ecosystem diversity, while metabarcoding identifies a greater number of individual species but shows a higher species dominance. The results suggest that microscopy provides a more complete picture of ecosystem diversity, while metabarcoding identifies a greater number of individual species but shows a higher species dominance.

Only 2% of taxa overlapped between the two methods, with Tabellaria flocculosa (Roth) Kützing and Psammothidium germainii (Manguin) Sabbe being the only two species that were concordant in both approaches. This extremely low level of overlap highlights significant discrepancies between the methods used in taxon identification, suggesting that they may capture different aspects of the community or have different sensitivities to particular species. As shown in Figure 4, the presence of T. flocculosa and P. germainii in both methods underscores the possible robustness or prevalence of these species in the studied ecosystem, despite the general lack of concordance among other taxa.

The apparent discrepancy between microscopy and metabarcoding results is probably not due to methodological flaws but reflects the unique characteristics of diatom communities in these environments. Our previous findings highlighted the prevalence of empty frustules from dead diatoms, detectable by microscopy but not by metabarcoding, and the presence of allochthonous propagules deposited through aeolian mechanisms. Each method offers distinct advantages: microscopy captures total diversity, including dead cells and empty frustules, while metabarcoding is more objective, focusing on viable organisms with intact DNA. Hence, both methods are considered complementary and essential for a comprehensive analysis, with microscopy providing historical composition and metabarcoding reflecting the active biological composition. Our approach accounts for specific challenges in studying cryoconite hole communities, including careful selection of primers and library preparation methods, while acknowledging the potential for further optimization. The dual approach offers a more nuanced understanding of these ecosystems, capturing both historical deposition and currently active community, which is crucial for accurate ecological assessments in these unique environments.

The use of metabarcoding to characterize diatoms in tropical ecosystems is relatively recent. This technique has primarily been applied in high-altitude rivers [12,24,35,64], while its application in tropical glaciers has been limited to bacterial characterization on the Cayambe and Sumaco Glacier [65]. The only study on diatoms in cryoconite holes of tropical glaciers was conducted by Chamorro et al. [10], who used optical microscopy to characterize these ecosystems and found a high diversity of taxa. One of the most representative and dominant species in the community was Psammothidium germainii. To confirm the optical identification of this species, we used scanning electron microscopy. (SEM) (Figure 5). However, the combined use of morphological and molecular characterization in the study of extreme habitats is essential for obtaining a comprehensive view of the diversity of diatoms present. Morphological characterization using optical microscopy allows for the identification of a wide range of taxa and the detection of both live and dead diatoms, as the valves of cells without chloroplasts can be observed, which would not be possible with metabarcoding, which only detects the DNA of living organisms. On the other hand, molecular characterization through metabarcoding is essential in these sediment-laden samples, where the observation of all species under the microscope is limited by the amount of material. This molecular approach enables the identification of a greater diversity of species, including those that are small or of low abundance, which might be overlooked by traditional methods.

For this reason, it is crucial to have a detailed and standardized protocol, such as the one described in this article, to guide future studies in these environments. This protocol not only ensures the proper preparation and processing of samples but also optimizes the integration of both methodologies to maximize the accuracy of species identification. Thus, it contributes to a more thorough and precise understanding of the ecological dynamics in these extreme ecosystems, providing a valuable reference for future research in tropical glaciers.

A key advancement in our methodology was the incorporation of metabarcoding, providing unprecedented insights into the diversity and composition of diatom communities within tropical glaciers. Metabarcoding, a high-throughput DNA-sequencing technique, allowed for the comprehensive analysis of genetic material in environmental samples, enabling the simultaneous identification of multiple diatom species [66,67].

By utilizing the 100 µm mesh prior to the digestion process, we optimized the extraction of genetic material, thereby improving the success of the metabarcoding analyses. This approach not only facilitated the identification of known diatom taxa but also uncovered potentially novel species, contributing to a more nuanced understanding of glacial ecosystems. The application of metabarcoding in our study aligns with contemporary trends in microbial ecology, providing a powerful tool for characterizing complex microbial communities in challenging environments [68,69].

The utilization of scanning electron microscopy (SEM) has served as a supplementary tool in specific instances, such as the identification of new species or the confirmation of uncertain species (Figure 5). It has not been employed as the primary means of characterizing an ecosystem. Nevertheless, its efficacy in observing marine diatoms [70] is well documented. SEM has proved to be useful for identifying highly complex, small species, or specific characteristics [71,72,73,74]. Additionally, SEM has found application as a supporting tool in other disciplines, including forensic science [75].

4. Conclusions

In conclusion, the characterization of diatom communities within cryoconites from tropical glaciers underscores the critical importance of integrating optical microscopy and metabarcoding for accurate species identification. This study presents a robust and accessible methodology for the collection, preparation, assembly, and identification of cryoconite hole samples from these unique ecosystems. However, it is important to note the limited number of studies specifically focused on diatom communities in cryoconite sediments. Many existing studies either lack detailed methodological descriptions or focus primarily on direct identification from collected samples, which poses a significant challenge to making meaningful comparisons with previous research.

The observed discrepancies between morphological and molecular inventories of diatom communities in cryoconite holes are likely due to the prevalence of empty frustules, which correspond to dead diatoms. While microscopy can detect these empty frustules, metabarcoding is unable to identify them due to the lack of intact genetic material. Microscopy provides a comprehensive view of all diversity, including both living cells and dead debris, while metabarcoding provides a more objective and unbiased approach by focusing only on viable organisms with intact DNA. This underscores the value of using both methods simultaneously: microscopy to capture total diversity (including both living and dead diatoms) and metabarcoding to examine the active biological community.

It is also important to note that barcoding studies in these ecosystems are still relatively new, and current reference databases may not include all species, limiting the completeness of molecular identification. Therefore, the use of a dual approach provides a more holistic understanding of diatom assemblages, combining insights into both current biological activity and the historical presence of diatoms in these unique environments.

Our protocol provides a refined approach to diatom characterization that, while providing valuable insights, complicates direct comparisons with previous studies. Consequently, while it is essential to situate our findings within the broader scientific context, the innovative methods employed in this research necessitate a more individualized interpretation of our results.