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Article

Impact of Blue Haslea spp. Blooms on Benthic Diatom and Bacterial Communities

by
Julie Seveno
1,2,
Andrzej Witkowski
3,
Ana Car
4,
Romain Gastineau
3,
Damien Sirjacobs
5,
Vincent Leignel
1 and
Jean-Luc Mouget
1,*
1
BIOSSE (Biologie des Organismes, Stress, Santé, Environnement) Laboratory, Le Mans University, 72000 Le Mans, France
2
Station de Recherches Sous-Marines et Océanographiques STARESO, 20260 Calvi, France
3
Institute of Marine and Environmental Sciences, University of Szczecin, Mickiewicza 16a PL, 70-383 Szczecin, Poland
4
Institute for Marine and Coastal Research, University of Dubrovnik, Kneza Damjana Jude 12, 20000 Dubrovnik, Croatia
5
InBioS–PhytoSYSTEMS Laboratory, University of Liège, B-4000 Liège, Belgium
*
Author to whom correspondence should be addressed.
Phycology 2024, 4(3), 465-507; https://doi.org/10.3390/phycology4030027
Submission received: 3 June 2024 / Revised: 29 August 2024 / Accepted: 1 September 2024 / Published: 11 September 2024
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Abstract

:
Climate change and nutrient enrichment are increasing the frequency of algal blooms, with sometimes significant impacts on coastal ecosystems. Haslea ostrearia blooms have been documented in oyster ponds and are not harmful, yet their effects in open environments remain underexplored. Marennine, a blue pigment produced by H. ostrearia, can display a range of biological properties in laboratory conditions, including antibacterial and allelopathic properties. Other blue Haslea species, forming blooms, synthesize bioactive marennine-like pigments. This study aims to understand if and how these blooms could affect the underlying community of microorganisms living in the biofilms. Morphological and molecular techniques were used to assess community dynamics during bloom events. Our findings indicate that blue Haslea blooms do not significantly alter the diatom or bacterial populations. However, they are paired with enhanced alpha diversity in the microbial communities. These observations suggest a complex interaction between bloom events and microbial dynamics. Additionally, this study expands our understanding of the bioactive properties of marennine-like pigments and their ecological roles, suggesting new avenues for biotechnological applications. This work underscores the importance of further research into the environmental and biological implications of blue Haslea blooms.

1. Introduction

Photoautotrophic benthic biofilms, or periphyton, are dynamic microenvironments at the base of benthic food webs, formed through a succession of well-defined steps [1,2]. Bacteria are usually the initial colonizers, followed by pennate diatoms and other microorganisms such as cyanobacteria, fungi and microfauna [3,4]. This establishes a hierarchy reflecting the organization of the populations based on their needs for light, nutrient concentration and grazing pressure [5,6]. Diatoms and heterotrophic bacteria form tight associations and can influence each other through chemotaxis [7,8]. Bacteria use quorum sensing to exchange information and synchronize their activity within the biofilm, meaning they communicate via the secretion and sensing of chemical signaling molecules. These auto-inducers can regulate gene expression within the bacterial community [9]. Diatoms interact through allelopathic compounds. Allelopathy was defined by Ricel in 1984 [10] as “any direct or indirect, harmful or beneficial effect by one plant (including microorganisms) on another through the release of chemicals that escape into the environment.” Benthic diatoms could be better acclimatized and more resistant than planktonic diatoms due to the high levels of chemical signaling molecules in the biofilm [11]. In biofilms, lipophilic allelopathic compounds are often more effective, as water-soluble compounds dissolve easily in the water column.
These phototrophic biofilms maintain a rich diversity both inter- and intra-species, with several genotypes of the same species found in a single biofilm [12]. Biofilm composition can quickly adapt to changes due to abiotic (environmental parameters) and biotic (microorganisms’ compounds and density) factors. Under favorable conditions, one diatom species can outcompete the rest of the community with a faster cell division rate, leading to bloom formation. Algal blooms have increased in frequency and intensity worldwide, often due to the increase in nutrients in water [13,14]; therefore, understanding bloom dynamics and estimating their effects on associated communities are valuable.
So far, blue Haslea blooms and their impacts have been studied only in oyster ponds. These blooms in semi-closed environments reach their maximum at the end of autumn (North Atlantic). Haslea ostrearia (Gaillon) Simonsen, the emblematic species of the genus, is known to produce a blue pigment called marennine. When oysters filter pond water to feed during a bloom of H. ostrearia, their gills turn green due to the fixation of marennine pigment in their tissue [15]. These green-gilled oysters have a socio-economical value in the Marennes Oléron area of France since they have been consumed for centuries and are considered more valuable than the non-pigmented oysters. Moreover, marennine has demonstrated species-dependent allelopathic properties under laboratory conditions [16,17,18]. Its antibacterial properties have also been confirmed, showing a response that varies depending on the strain and on the species tested.
Previous studies in Atlantic oyster ponds during H. ostrearia blooms have shown a diminution of the density of Skeletonema costatum (Greville) Cleve populations [19,20]. Regarding the antibacterial effect against the genus Vibrio, the inhibition was also species and strain dependent [21].
Blue Haslea blooms also occur in open environments (Figure 1), for instance, in the Mediterranean Sea, where H. ostrearia and H. provincialis Gastineau, Hansen & Mouget were identified. These blooms happen in spring, after the phytoplankton bloom [22]. The biological activities of the pigment produced by H. provincialis have not yet been tested. However, it can be hypothesized that all marennine-like pigments possess bioactive properties similar to marennine, as evidenced in H. karadagensis Davidovich, Gastineau & Mouget [23] and in H. nusantara Mouget, Gastineau & Syakti [24]. In natural, open environments, the dynamics of blue Haslea blooms have recently been documented, but their possible impacts remain to be studied. Indeed, the biological effects of marennine-like pigments might differ in an open environment compared to under laboratory conditions or in oyster ponds. In these closed or semi-closed environments, marennine-like pigments can accumulate and reach a threshold at which impacts on organisms are noticeable, whereas, in open environments, hydrophilic pigments could be rapidly diluted in the water column. Moreover, interactions between organisms within biofilms are very diverse and complex [25,26,27,28], which could modify biological responses, thus differing from the straightforward biological effects observed in laboratory conditions. The objective of this study was to estimate the impact of blue Haslea blooms on the main diatom and bacterial communities of the underlying photoautotrophic biofilm in two different areas of the western Mediterranean Sea. These two distinct sites were selected to measure the impact of algal blooms in areas with varying environmental characteristics, allowing for a comparison and validation of the observed effects.

2. Materials and Methods

2.1. Study Area

Two areas in the Mediterranean Sea were chosen for the significant presence of blue Haslea blooms to study and compare the impact of these blooms on biofilm communities. Both of these areas are considered oligotrophic and are referred to as non-polluted sites. The first area was located in Calvi Bay on the northwestern coast of Corsica Island (France), within the Ligurian Sea. Sampling was conducted on 12 and 13 April 2019, and from 25 to 29 March 2021. Along the Corsican coast, in 2019, two sampling sites were chosen, one with low-intensity blue Halslea patches (Port site) and one where Haslea bloom was the most developed (Madrepore site). In 2021, the bloom was not as developed on the Madrepore site, so a third site was added where the bloom was the most developed (Oscelluccia site). In Corsica, sampling was carried out by scuba diving from the STARESO marine station. The second area was along the Croatian coast, between Hvar Island and the Dubrovnik coast, with sampling from 21 to 26 July 2020. Along the Croatian coast, three sites were chosen, and sampling was performed by snorkeling at these three sites (Table 1, Figure 2). As they were different geographical areas, environmental parameters were recorded at each site; see Materials and Methods [22] for a full description.
Two different substrates were selected, the brown algae Padina Adanson sp. thalli and turf, and the same substrates without blue Haslea blooms were used as controls. At each site, three quadrats were placed in the bluest zones and three in the control zones (where no blue Haslea patches were visible). Samples taken in bloom conditions were a few meters (maximum 3 meters) away from the control ones. In each quadrat, one 15 mL tube was filled with Padina sp. Thallus, and, in Corsica, a second 15 mL tube was filled with turf. Turf was not sampled in Croatia because the bloom was only observed on Padina sp. thalli. For diatom observations, tubes were preserved in 4% formaldehyde at room temperature until further processing. In the same quadrat, when sampling for the bacterial community was conducted, one 50 mL tube was filled with Padina sp. and another with turf. These tubes were preserved in 96% ethanol at -80 °C until DNA extraction. In each quadrat and for both substrates, a fresh fraction was taken to measure the blue Haslea spp. density to compare it with the rest of the total diatom community.

2.2. Samples Process for Diatom Analysis

To remove the organic matter, samples were boiled in hydrogen peroxide (30%) for 24 h. They were then washed with deionized water. Ten drops of the suspension were air-dried onto coverslips which were subsequently mounted onto glass slides using Naphrax® glue (Brunel Microscopes Ltd., Wiltshire, UK) [29,30].
The slides were observed under a light microscope (LM), the Zeiss Axio Scope A1, equipped with a ×100 Plan Apochromatic immersion objective with differential interference contrast (DIC) (Carl Zeiss Ltd., Oberkochen, Germany). Pictures were taken using a Zeiss Axio Cam ICc5 camera (Carl Zeiss Ltd., Oberkochen, Germany). With the LM, a minimum of 300 diatom valves were counted and identified [31,32,33,34]. Haslea cells, which have very fine valves that are difficult to discern under LM, were not counted to better visualize variation in the diatom community.
Scanning Electron Microscopy (SEM) was used to verify the taxonomy of some species. Samples were pipetted onto polycarbonate membrane filters (Whatman Nuclepore with 1 µm pores), then mounted onto aluminium stubs and sputter-coated with 20 nm of gold or gold–palladium alloy using a Q150T coater from Quorum Technologies (Laughton, UK). Observations were performed using the ultra-high-resolution scanning microscope SU8020 from Hitachi (Tokyo, Japan), provided by the University of Szczecin (Szczecin, Poland).

2.3. Bacterial DNA Extraction, PCR Amplification and Data Processing

DNA extraction was performed using the Power Soil DNA Isolation Kit (Mo Bio, Germantown, MD, USA) according to the manufacturer’s instructions. DNA quality and quantity were verified using a Nanodrop 2000 (Thermo Fisher Scientific, Wilmington, DE, USA) and adjusted to 2.0 × 10−2 g/L. The V3 and V4 hypervariable regions of 16S rDNA were amplified by PCR. Primer design and libraries were prepared by Azenta, Leipzig, Germany (16S-EZ), which then proceeded to Illumina-MiSeq sequencing (250 bp paired end,). Cutadapt software (v1.9.1) was used to merge each read pair. Qiime software (1.9.1) [35] was employed to verify the quality of the sequencing and remove chimeric sequences. Sequences with more than 97% similarity to OTUs were considered the same OTU. The total number of reads per sample ranged from 14,704 to 51,275. Species annotation was performed by comparing OTUs to the Greengenes database using the RDP classifier Bayesian algorithm on Qiime software (version 1.8.0) [36].

2.4. Statistical Analysis

The graphics were produced using the ggplot2 package in the R software (https://www.r-project.org/, version 4.0.3). A Hellinger transformation was applied to total bacterial abundance [37]. In the analysis, either the RA or Hellinger abundance (HA) was used. In taxonomy abundance bar plots, ‘Other’ represents phylum-level taxa with a relative abundance (RA) greater than 1%. The Hopkins statistic was employed to verify the possibility of clustering. The K-means method was used for clustering on the PCA. Alpha diversity was estimated using Shannon and Simpson indexes via the vegan package in R [38,39]. The Shannon index allows comparison between samples, and the Simpson index can also be compared with indexes from other studies. T-tests were performed to verify the statistical significance between conditions. Beta diversity was measured using unweighted and weighted UniFrac [40,41]. Non-Metric Multidimensional Scaling (NMDS) was conducted on the beta diversity distance matrix to visualize sample clustering using the vegan package in the R software. A permutational multivariate analysis of variance (PERMANOVA) (Adonis3, package GUniFrac) based on Bray–Curtis dissimilarity with 999 permutations was performed to verify the results of clustering.

3. Results

3.1. Environmental Parameters

When samples were collected, the seawater temperature was between 14.5 °C and 15.0 °C in Corsica and 23.6 °C in Croatia. The average salinity was 37.9 psu in Corsica and 38.7 psu in Croatia. During sample collection in Corsica, NO3- ranged from 0.10 µM to 0.58 µM; PO43- ranged from 0.01 μM to 0.04 μM; SiO44- ranged from 0.51 μM to 0.65 μM. During sample collection in Croatia, NO3- ranged from 0.22 µM to 0.27 µM; PO43- ranged from 0.11 μM to 0.36 μM; SiO44- ranged from 2.6 μM to 6.1 μM. For a detailed analysis of the interaction between environmental parameters and the presence of Haslea bloom, see [22].

3.2. Diatom Community

In the 43 samples from the Mediterranean Sea (Corsican and Croatian samples), 95 genera of diatoms were identified in total, with 16 to 38 different genera per sample. Haslea species H. provincialis and H. ostrearia accounted for 46% of the identified species. The five other most abundant genera, grouping 70% of the total identified species, were Cocconeis (27%), Mastogloia (23%), Navicula (12%), Nitzschia (5%) and Grammatophora (3%). In the Corsican samples, Cocconeis and Mastogloia were the predominant genera, whereas, in Croatia, Mastogloia was the most abundant (Figure 3). Cocconeis was the only genus present in every sample. Mastogloia was the most diverse genus, with 42 different species identified. Among the samples, the top ten most abundant species were Cocconeis scutellum Ehrenberg (15%), Cocconeis molesta Kützing (5.5%), Cocconeis diaphana W. Smith (4.9%), Mastogloia cuneata (F.Meister) Simonsen (4%), Mastogloia fimbriata (T.Brightwell) Grunow (3.7%), Toxarium undulatum Bailey (3.1%), Navicula cf. normalis Hustedt (2.9%), Mastogloia binotata (Grunow) Cleve (2.4%), Navicula sp. (2.3%) and Mastogloia crucicula (Grunow) Cleve (1.7%). All the diatom community compositions and corresponding micrographs are presented in the Appendix A, Figure A1, Figure A2, Figure A3, Figure A4, Figure A5, Figure A6, Figure A7, Figure A8, Figure A9, Figure A10, Figure A11, Figure A12, Figure A13, Figure A14, Figure A15, Figure A16, Figure A17, Figure A18, Figure A19, Figure A20, Figure A21, Figure A22, Figure A23, Figure A24, Figure A25, Figure A26 and Figure A27.
Variations in the diatom community according to sampling location were observed; Amphora spp., Berkeleya spp., Licmophora spp. and Nitzschia spp. had higher relative abundance (RA) in Croatia, while Ardissonea spp., Cocconeis spp., Grammatophora spp. and Surirella spp. had higher RA in Corsica (Figure 3). However, no significant differences were found between the alpha diversity indices (Shannon and Simpson) at the two locations.
Substrates have also influenced the structure of the diatom community, with more Rhopalodia spp. on turf and more Achnanthes spp. and Berkeleya spp. on Padina sp. Alpha diversity indices (Shannon and Simpson) were not significantly different between the epiphytic diatom communities of turf and Padina sp.
Differences in diatom community composition between control and blue Haslea blooms were noted. Cocconeis spp. accounted for an average of 31% of the diatom community in control samples compared with an average of 20% in blue samples. This difference was significant, as demonstrated by a t-test on either RA (p-value = 0.013) or the absolute abundance standardized by Hellinger method (p-value = 0.04). At the species level, C. scutellum and C. molesta were less present in blue Haslea bloom samples, with respective averages of 18% and 6% RA compared with 10% and 3% RA in control zones (C. scutellum t-test p-value = 0.018; C. molesta t-test p-value = 0.039). Inversely, Diploneis spp. (not identified to the species level) had a significant increase (1.9%) in RA in the presence of Haslea bloom (RA p-value = 0.04; HA p-value = 0.009). Despite Mastogloia being a species-rich genus, no significant changes were observed in samples with or without blue Haslea blooms at the genus or species level.
The alpha diversity of samples taken during blue Haslea blooms was significantly higher compared with control samples (Supplementary Material, Figure S1).
Samples could be clustered according to their diatom distribution (Figure 4a). As illustrated in Figure 4a, Croatian samples seem to be regrouped, but they are not forming one cluster. There was a clear opposition between Mastogloia and Cocconeis, with the presence of Mastogloia correlating with samples from Croatia, where it was the main genus. Cocconeis spp. were more prevalent in Corsican samples. Navicula and Grammatophora also played significant roles in terms of RA and structured the population. The PCA shows that the presence or absence of blue Haslea sp. was not the main component structuring the diatom community (Figure 4b); blue and control samples are mixed in the different groups.

3.3. Bacterial Community

The bacterial communities of 24 samples taken from Calvi Bay in 2021 were investigated across two different sites and two substrates (Padina sp. and turf), with, in each case, samples taken within the bloom and control samples, with three replicates per condition. The main orders, representing more than 10% of the bacterial community in at least one sample, were Acidobacteriales, Alteromonadales, Bacillales, Clostridiales, Flavobacteriales, HTCC2188, Legionellales, Rhizobiales, Rhodobacterales, Rickettsiales, Sphingomonadales and Vibrionales (Figure 5). About 31.8% of bacteria were unidentified and unassigned to a bacterial order.
The two sites along the Revelleta Cape, Port and Oscelluccia, were 500 m apart. Only the bacterial order Desulfobacterales was more present in the Port site (p-value = 0.02). Substrate type also influenced the bacterial community; the RA of Bacillales and Burkholderiales was higher on Padina sp., in contrast to Cerasicoccales, Marinicellales, Thiotrichales, Thiohalorhabdales and Verrucomicrobiales, which had a higher RA on turf. Nevertheless, the alpha diversity indices—Shannon and Simpson—did not differ significantly between sites or substrates.
The dominant orders in blue Padina sp. samples were Rhodobacterales (22.6%) and Flavobacteriales (10.5%), whereas, in control Padina samples, Rhodobacterales (15.3%) and Rhizobiales (14.1%) were dominant (Figure 5a). On turf, in both bloom and control samples, Rhodobacterales (27.8% blue; 21.1% control) and Flavobacteriales (12.5% blue; 11.9% control) were the main orders (Figure 5b).
In bloom samples, the orders Alteromonadales, Flavobacteriales, Rhodobacterales and Sphingomonadales exhibited a significantly higher relative abundance compared to other orders. Although unidentified genera represented 85% of the total bacterial community, variations in bacterial distribution were noted. The genera Dinoroseobacter and Kordia were more prevalent in bloom conditions, whereas Hyphomicrobium and Leptolyngbya were less abundant in bloom samples. Anaerococcus was only identified in bloom samples. In blooming conditions, Shannon and Simpson indices had mean values of, respectively, 7.44 and 0.97 compared to 6.46 and 0.89 in control conditions. The alpha diversity of bacterial communities was significantly higher in bloom samples (t-test, Shannon p-value = 0.029, Simpson p-value = 0.04; see Supplementary Material, Figure S2). An NMDS analysis was performed to verify how the bacterial communities of each sample were distributed. Based on the second axis, Haslea bloom samples seemed to form a group apart from the control samples (Figure 6). However, PERMANOVA showed that the trend was not significant.

4. Discussion

This study provides new insights into the ecological dynamics of benthic periphyton communities in the western Mediterranean Sea during blue Haslea spp. blooms. This topic has emerged thanks to previous laboratory findings that highlighted the allelopathic and antibacterial properties of marennine-like pigments produced by blue Haslea cells [17,18,21,23]. Our research aimed to assess whether these effects manifest similarly in natural open environments in the western Mediterranean Sea, and to estimate their ecological importance, if any. The results did not indicate critical shifts in the communities. However, notable variations were observed, with a marked increase in species diversity in samples collected during bloom events. In particular, diatom and bacteria diversity was higher in samples collected during blue Haslea blooms.

4.1. Diatom Community Associated within the Biofilm

Diatoms serve as effective environmental indicators as the community can evolve rapidly in response to changing environmental parameters [42]. Differences between locations can be attributed to geographical distance and to seasonal fluctuations. For instance, samples from the Ligurian Sea were collected in March, whereas those from the Adriatic Sea were collected in July. Epiphytic diatom composition has been proven to be host related [43,44]. The study by Car et al. (2019) [29] analyzed the epiphytic diatom community on Padina sp. and had analogous results. Mastogloia spp. represented an average of 27% of the total diatom community in their samples, compared to 29% in our Croatian samples and 23% generally. Additionally, the study found that Cocconeis, Hyalosynedra, Navicula and Nitzschia each accounted for more than 5% of the total diatom community, analogous to our findings in Croatian samples.
When comparing bloom samples with control ones, a diminution of the RA of C. scutellum and C. molesta and an increase in the RA of Diploneis spp. were observed. Cocconeis spp. are epiphytes strongly attached to large-thallus algae and phanerogams such as Caulerpa spp., Posidonia oceanica (Linnaeus) Delile, Zostera marina Linnaeus and Nymphea alba Linnaeus [45,46,47,48]. Conversely Diploneis spp. is moderately attached and often found on periphyton, seagrasses and sponges [49,50]. The study of blue Haslea bloom effects showed a genus- or even species-dependent response. For instance, Cocconeis and Diploneis were affected in opposite ways by blue Haslea blooms despite occupying similar ecological niches.
In both areas, in blue Haslea bloom samples, an increase in diatom diversity was observed. The calculation of diversity indexes is based on the number of species and their distribution. A decrease in the abundance of the dominant species, Cocconeis spp., can lead to a more homogeneous distribution of species abundance in the samples, therefore improving alpha diversity overall.
During blue Haslea blooms in semi-closed and shallow environments like oyster ponds, the marennine pigment can accumulate in the water column [51], reaching high concentrations of up to 5 × 10−3 g/L [52]. This accumulation has been shown to potentially exert allelopathic effects on planktonic diatom communities. Previous studies [19,53] have noted poorer flora in oyster ponds during blue Haslea blooms, with H. ostrearia representing a high percentage of the diatom community. The composition and role of marennine in relation to H. ostrearia are not yet fully understood. As producing this pigment has an energetic cost, it must provide environmental benefits to the cells. Given that marennine is a hydrophilic pigment, its effects may primarily target planktonic diatoms. This dynamic could be particularly relevant in shallow oyster beds, where the pigment is less likely to dilute, and its inhibitory effect on planktonic organisms could be sustained with less energy expenditure [54].
In the Mediterranean samples, marennine and marennine-like pigments were present intracellularly. The extracellular form was not detectable in seawater, likely due to a rapid dilution in the water column. As a result, any potential allelopathic effects are likely limited to the microenvironment around the cells. Blue Haslea cells developed on top of the biofilm, where they had better access to light, creating a shading effect that potentially affected the growth rates of other diatoms and phototrophic microorganisms. This shading effect could allow Haslea to outcompete other diatoms [17].
Moreover, in these oligotrophic environments, access to nutrients could also be part of the competition between blue Haslea and other diatoms. During blooms, cells have a fast reproduction rate and silica may become a limiting nutrient [55,56,57]. In our samples, some diatoms appeared with dissolved valves (Figure A27), suggesting that benthic blooms of Haslea might utilize a significant portion of the dissolved silica. Taken together, these observations suggest that blue Haslea spp. are opportunistic species, outcompeting the diatom community by creating shading effects and reducing access to light and nutrients, especially silica, although this impact appears to be limited.

4.2. Bacteria Associated with the Biofilm

Bacteria play a key role for microalgae, especially during blooms, as they can quickly remineralize the organic matter and provide essential co-factors like B12 vitamins. The major bacterial orders found in our samples were Flavobacteriales, Rhizobiales and Rhodobacterales. Flavobacteriales commonly associated with phytoplankton blooms efficiently degrade high-molecular-weight macromolecules from microalgae [58,59,60], facilitating carbon remineralization [61]. Rhizobiales bacteria are commonly associated with plant roots due to their capability to fix nitrogen [62] and have been associated with mollusk microbiomes [63,64] and are epiphytes on various algae [65]. Rhodobacterales are an important part of the diatom microbiome [66] as 90% of their species are able to fully synthesize the essential vitamin B12 [65]. Other significant orders encountered in samples belong to the phyla Acidobacteriota (Acidobacteriales and Acidomicrobiales), Alpha-proteobacteria (Rickettsiales and Sphingomonadales), Gamma-proteobacteria (Alteromonadales, HTCC2188, Legionellales and Vibrionales) and Firmicutes (Bacillales and Clostridiales), which are frequently associated with algae [67]. The order HTCC2188 regroups marine bacteria from oligotrophic seas such as the Mediterranean Sea [68]. Acidobacteriales have also been reported in planktonic and soil environments in the Mediterranean Sea [69]. The abundance of Vibrionales has been shown to increase in diatom-dominated blooms [70].
The same bacterial families (Rhodobacteraceae, Flavobacteriaceae, Saprospiraceae and Verrucomicrobiaceae) were identified in our samples, as well as in those collected from the macroalgae Cystoseira compressa (Esper) Gerloff & Nizamuddin in the Mediterranean Sea [71]. In another study, on the prokaryotic community of Taonia sp. (Dictyotaceae family) sampled on the French Mediterranean coast, the Saprospiraceae family was dominant [72]; however, it was not observed in our samples. The substrate type could explain this discrepancy as it also influenced the bacterial community in our samples; Bacillales (Firmicutes) and Burkholderiales (Beta-Proteobacteria) were exclusively found on Padina sp. Furthermore, these phyla and orders have already been encountered as Padina epiphytes along the Tunisian coast [73].
Significant differences were observed within bacterial communities depending on the presence or lack of Haslea bloom in both locations. Typically, in biofilm, aerobic bacteria are closer to the boundary layer, while anaerobic bacteria reside at the bottom of it. Anaerococcus, an anaerobic genus [74], was uniquely identified in bloom samples, aligning with the expected oxygen gradient in biofilm, which can be even more important during an active bloom [75,76]. Dinoroseobacter, an aerobic genus with photosynthetic pigments that have been previously associated with phototrophic dinoflagellate blooms [77], was also found in higher proportions during Haslea blooms. The genus Kordia, more abundant in bloom samples, can act as an algaecide for certain diatoms but showed variable effects depending on the species [78], suggesting a potential to influence bloom dynamics over time.
Two bacterial genera were found in lower proportions during bloom conditions—Leptolyngbya and Hyphomicrobium—accounting for only a minor fraction of the community, 0.1% and 0.02%, respectively. The genus Hyphomicrobium can denitrify biofilm compounds in a salt-water treatment facility, and some species are known to be methylotrophic [79,80]. The cyanobacteria Leptolyngbya, which have a cosmopolitan distribution, are halotolerant and able to grow in extreme conditions. They were tested in agro-industrial wastewater treatments due to their ability to recycle nitrogen and phosphorus in high concentrations [81]. Some species of the Leptolyngbya genus are known as worldwide coral pathogen agents [82].
Comparing the bacterial community associated with a bloom of blue Haslea spp. with the bacterial community of H. ostrearia cells from the French Atlantic coast, whether newly isolated [83] or from cells of a culture collection [84], reveals consistent associations with orders like Flavobacteriales, Oceanospirillales, Pseudomonadales, Rhodobacterales, Rickettsiales and Sphingomonadales. Some orders and genera associated with H. ostrearia in laboratory conditions were not detected in our samples or in newly isolated strains. This discrepancy may be due to differences in sampling locations or seasonal variations. Moreover, in the present work, H. provincialis accounted for the majority of bloom events, suggesting that bacterial communities may vary depending on the blue Haslea species present. Additionally, it has been observed that, after three months in laboratory culture, there is a noticeable shift in the bacterial community associated with H. ostrearia [85]. These observed changes with time in bacteria communities associated with blue Haslea cells are still scarce, but they reinforce the need for more in-depth studies to identify the bacterial communities in blue Haslea phycospheres and compare their evolution from in situ conditions to long-lasting cultures. This could reveal essential information, both for strain maintenance in the laboratory and for culture upscaling for industrial application. Indeed, the interactions between bacteria and diatoms are species specific; therefore, each diatom species has a unique holobiont [86].

5. Conclusions

Photoautotrophic benthic biofilms are complex structures composed of numerous interdependent microorganisms influenced by environmental conditions [87]. This microenvironment has its own dynamics distinct from the water column. Biofilms are characterized by a high density of microorganisms and a corresponding high density of signaling chemical compounds, making benthic microorganisms more resistant to allelopathic compounds [11,88,89].
Our findings indicated that the presence of blue Haslea spp. blooms did not have significant negative effects on the biofilm community. On the contrary, an increase in diversity within diatom and bacterial communities was observed. To date, there are no examples of pathology associated with the consumption of green oysters. Therefore, these benthic blooms are not classified as harmful.
Overall, this study underscores the complexity of interactions within photoautotrophic benthic biofilms and highlights that Haslea blooms are not harmful to the environment despite the many biological activities of marennine-like pigments. Instead, blue Haslea blooms appear to positively influence the diversity of diatoms and bacteria by decreasing the relative abundance of dominant diatom species.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/phycology4030027/s1, Figure S1. Simpson and Shannon indexes for the diatom community. T-test was performed to verify the significance in between samples; there was a significant change in the diversity in between control and blue zones (p-value < 0.05). Figure S2. Alpha diversity was measured for the bacterial population, and a t-test was performed to verify the significance in between conditions in blank or blue zones. (a) Shannon index p-value = 0.029; (b) Simpson index t-test p-value = 0.04.

Author Contributions

Conceptualization, J.S. and J.-L.M.; methodology, A.C., D.S. and J.S.; formal analysis, A.W. and R.G.; data curation, A.W.; writing—original draft preparation, J.S.; writing—review and editing, J.S.; supervision, V.L. and J.-L.M.; visualization, V.L., A.W. and R.G.; funding acquisition, J.-L.M. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Horizon 2020 Research and Innovation Program GHaNA (“The Genus Haslea, New marine resources for blue biotechnology and Aquaculture”, grant agreement no. [734708/GHANA/H2020-MSCA-RISE-2016], J.-L.M.). Observations along the Adriatic Sea coast were partly funded by the Croatian Science Foundation under project IP-2019-04-9043 (DiVMAd).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in the study are included in the article/supplementary material, further inquiries can be directed to the corresponding author.

Acknowledgments

Thanks to the STARESO staff for diving support and the availability of data on Calvi Bay from the program STARECAPMED (Station of Reference and Research on Change of Local and Global Anthropogenic Pressures on Mediterranean Ecosystem Drifts, supported by Territorial Collectivity of Corsica and the Rhone-Mediterranean and Corsica Water Agency). We thank A.P., A.S. and A.W. for their statistical advice. We thank Zoran Jurić from the University of Dubrovnik for organizing the excursion along the Adriatic coast.

Conflicts of Interest

The authors declare no conflicts of interest.

Appendix A

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Figure A1. (1.1) Coscinodiscus Ehrenberg sp., (1.2) Thalassiosira Cleve sp., (1.3) Thalassiosira oestrupii (Ostenfeld) Proshkina-Lavrenko, (1.4) Actinoptychus sp1., (1.5) Actinoptychus Ehrenberg sp2., (1.6) Triceratium pentacrinus (Ehrenberg) Wallich, (1.7) Trigonium diaphanum A.Mann.
Figure A1. (1.1) Coscinodiscus Ehrenberg sp., (1.2) Thalassiosira Cleve sp., (1.3) Thalassiosira oestrupii (Ostenfeld) Proshkina-Lavrenko, (1.4) Actinoptychus sp1., (1.5) Actinoptychus Ehrenberg sp2., (1.6) Triceratium pentacrinus (Ehrenberg) Wallich, (1.7) Trigonium diaphanum A.Mann.
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Figure A2. (2.1) Auliscus Ehrenberg sp., (2.2,2.2bis) Lampriscus shadboltianum (Greville) H.Peragallo & M.Peragallo, (2.3) Biddulphia tridentula Ehrenberg, (2.4) Cymatosira lorenziana Grunow, (2.5) Cyclostephanos Round sp., (2.6) Odontella Round sp., (2.7) Biddulphia biddulphiana (J.E.Smith) Boyer.
Figure A2. (2.1) Auliscus Ehrenberg sp., (2.2,2.2bis) Lampriscus shadboltianum (Greville) H.Peragallo & M.Peragallo, (2.3) Biddulphia tridentula Ehrenberg, (2.4) Cymatosira lorenziana Grunow, (2.5) Cyclostephanos Round sp., (2.6) Odontella Round sp., (2.7) Biddulphia biddulphiana (J.E.Smith) Boyer.
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Figure A3. (3.1) Skeletonema Greville sp, (3.2) Lampriscus shadboltianum.
Figure A3. (3.1) Skeletonema Greville sp, (3.2) Lampriscus shadboltianum.
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Figure A4. (4.1) Grammatophora Ehrenberg sp., (4.2) Fragilaria Lyngbye sp1., (4.3) Hendeyella M.P.Ashworth, Witkowski & CL.Li sp., (4.4) Licmophora remulus Grunow, (4.5) Licmophora C.Agardh sp1., (4.6) L. abbreviate C.Agardh, (4.7) Licmophora sp2., (4.8) Licmophora sp3., (4.9) Licmophora arcuata Car & Herwig.
Figure A4. (4.1) Grammatophora Ehrenberg sp., (4.2) Fragilaria Lyngbye sp1., (4.3) Hendeyella M.P.Ashworth, Witkowski & CL.Li sp., (4.4) Licmophora remulus Grunow, (4.5) Licmophora C.Agardh sp1., (4.6) L. abbreviate C.Agardh, (4.7) Licmophora sp2., (4.8) Licmophora sp3., (4.9) Licmophora arcuata Car & Herwig.
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Figure A5. (5.1) Divergita toxoneides (Castracane) Theriot, (5.2–5.4) Neosynedra provincialis (Grunow) D.M.Williams & Round, (5.5) Tabularia fasciculate (C.Agardh) D.M.Williams & Round, (5.6,5.7) Hyalosynedra D.M.Williams & F.E.Round sp., (5.8) Thalassionema Grunow sp1., (5.9) Thalassionema sp2., (5.10) Diatoma cf. vulgaris Bory, (5.11) Hyalosira Kützing sp., 11 Podocystis adriatica (Kützing) Ralfs, (5.12) Podocystis spathulata (Shadbolt) Van Heurck.
Figure A5. (5.1) Divergita toxoneides (Castracane) Theriot, (5.2–5.4) Neosynedra provincialis (Grunow) D.M.Williams & Round, (5.5) Tabularia fasciculate (C.Agardh) D.M.Williams & Round, (5.6,5.7) Hyalosynedra D.M.Williams & F.E.Round sp., (5.8) Thalassionema Grunow sp1., (5.9) Thalassionema sp2., (5.10) Diatoma cf. vulgaris Bory, (5.11) Hyalosira Kützing sp., 11 Podocystis adriatica (Kützing) Ralfs, (5.12) Podocystis spathulata (Shadbolt) Van Heurck.
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Figure A6. (6.1) Toxarium undulatum Bailey, (6.2) Toxarium hennedyanum (W.Gregory) Pelletan, (6.3) Tessella interrupta Ehrenberg, (6.4) Pteroncola R.W.Holmes & D.A.Croll sp., (6.5) Ardissonea crystalline (C.Agardh) Grunow, (6.6) A. formosa (Hantzsch) Grunow, (6.7) Cyclophora tenuis Castracane.
Figure A6. (6.1) Toxarium undulatum Bailey, (6.2) Toxarium hennedyanum (W.Gregory) Pelletan, (6.3) Tessella interrupta Ehrenberg, (6.4) Pteroncola R.W.Holmes & D.A.Croll sp., (6.5) Ardissonea crystalline (C.Agardh) Grunow, (6.6) A. formosa (Hantzsch) Grunow, (6.7) Cyclophora tenuis Castracane.
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Figure A7. (7.1) Licmophora sp., (7.2) Tabularia fasciculate (C.Agardh) D.M.Williams & Round, (7.3) Pteroncola sp., (7.4) Hyalosynedra sp., (7.5) Hyalosynedra sublaevigata Álvarez-Blanco & S.Blanco, (7.6) Protoraphis R.Simonsen sp., (7.7) Cyclophora tenuis Castracane, (7.8) Falcula M.Voight sp.
Figure A7. (7.1) Licmophora sp., (7.2) Tabularia fasciculate (C.Agardh) D.M.Williams & Round, (7.3) Pteroncola sp., (7.4) Hyalosynedra sp., (7.5) Hyalosynedra sublaevigata Álvarez-Blanco & S.Blanco, (7.6) Protoraphis R.Simonsen sp., (7.7) Cyclophora tenuis Castracane, (7.8) Falcula M.Voight sp.
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Figure A8. (8.1) Delphineis australis (P.Petit) Tsuy.Watanabe, Ji.Tanaka, G.Reid, Kumada & Nagumo, (8.2,8.2bis) Delphineis surirella (Ehrenberg) G.W.Andrews, (8.3) Striatella unipunctata (Lyngbye) C.Agardh, (8.4–8.6) Plagiogramma Greville spp.
Figure A8. (8.1) Delphineis australis (P.Petit) Tsuy.Watanabe, Ji.Tanaka, G.Reid, Kumada & Nagumo, (8.2,8.2bis) Delphineis surirella (Ehrenberg) G.W.Andrews, (8.3) Striatella unipunctata (Lyngbye) C.Agardh, (8.4–8.6) Plagiogramma Greville spp.
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Figure A9. (9.1,9.2,9.2bis) Achnanthes armillaris (O.F.Müller) Guiry, (9.3) Achnanthes parvula Kützing, (9.4) Cocconeiopsis Witkowski, Lange-Bertalot & Metzeltin sp., (9.5) Planothidium Round & Bukhtiyarova sp1., (9.6) Planothidium sp2., (9.7) Planothidium sp3.
Figure A9. (9.1,9.2,9.2bis) Achnanthes armillaris (O.F.Müller) Guiry, (9.3) Achnanthes parvula Kützing, (9.4) Cocconeiopsis Witkowski, Lange-Bertalot & Metzeltin sp., (9.5) Planothidium Round & Bukhtiyarova sp1., (9.6) Planothidium sp2., (9.7) Planothidium sp3.
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Figure A10. (10.1) Amphicocconeis M.De Stefano & D.Marino sp., (10.2) Cocconeis britannica Naegeli ex Kützing, (10.3) C. diaphana W.Smith, (10.4) Cocconeis sp1., (10.5) Cocconeis sp2., (10.6) C. neothumensis Krammer, (10.7,10.8) C. krammeri Lange-Bertalot & Metzeltin, (10.9,10.10) Cocconeis sp3., (10.11) C. molesta Kützing, (10.12,10.12bis) C. scutellum Ehrenberg, (10.13) C. scutellum var. posidonniae M.De Stefano, D.Marino & L.Mazzella.
Figure A10. (10.1) Amphicocconeis M.De Stefano & D.Marino sp., (10.2) Cocconeis britannica Naegeli ex Kützing, (10.3) C. diaphana W.Smith, (10.4) Cocconeis sp1., (10.5) Cocconeis sp2., (10.6) C. neothumensis Krammer, (10.7,10.8) C. krammeri Lange-Bertalot & Metzeltin, (10.9,10.10) Cocconeis sp3., (10.11) C. molesta Kützing, (10.12,10.12bis) C. scutellum Ehrenberg, (10.13) C. scutellum var. posidonniae M.De Stefano, D.Marino & L.Mazzella.
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Figure A11. (11.1) Cocconeis britanica, (11.2) C. caulerpacola Witkowski, Car & Dobosz, (11.3) C. peltoides Hustedt, (11.4) C. molesta, (11.5) Cocconeis molesta var. crucifera Grunow, (11.6) C. stauroneiformis H.Okuno, (11.7 C. scutellum var. posidoniae (&pores details), (11.8) C. scutellum, (11.9) C. scutellum douple-pored.
Figure A11. (11.1) Cocconeis britanica, (11.2) C. caulerpacola Witkowski, Car & Dobosz, (11.3) C. peltoides Hustedt, (11.4) C. molesta, (11.5) Cocconeis molesta var. crucifera Grunow, (11.6) C. stauroneiformis H.Okuno, (11.7 C. scutellum var. posidoniae (&pores details), (11.8) C. scutellum, (11.9) C. scutellum douple-pored.
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Figure A12. (12.1) Berkeleya cf. hyalina (Round & M.E.Brooks) E.J.Cox, (12.2) B. scopulorum (Brébisson ex Kützing) E.J.Cox, (12.3) Berkeleya sp., (12.4–12.6) B. fennica Juhlin-Dannfelt, (12.7) Gomphonema Ehrenberg sp., (12.8) Gomphonemopsis Medlin sp., (12.9) Rhoicosphenia Grunow sp.
Figure A12. (12.1) Berkeleya cf. hyalina (Round & M.E.Brooks) E.J.Cox, (12.2) B. scopulorum (Brébisson ex Kützing) E.J.Cox, (12.3) Berkeleya sp., (12.4–12.6) B. fennica Juhlin-Dannfelt, (12.7) Gomphonema Ehrenberg sp., (12.8) Gomphonemopsis Medlin sp., (12.9) Rhoicosphenia Grunow sp.
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Figure A13. (13.1) Tetramphora decussata (Grunow) Stepanek & Kociolek, (13.2) Amphora immarginata Nagumo, (13.3) Halamphora hyalina (Kützing) Rimet & R.Jahn, (13.4) A. pseudohylina Simonsen, (13.5,13.6) A. helenensis Giffen, (13.7,13.8) A. wisei (M.M.Salah) Simonsen, (13.9) A. marina W.Smith, (13.10). Amphora sp1., (13.11,13.12) Seminavis cf. robusta D.B.Danielidis & D.G.Mann, (13.13) Seminavis sp1., (13.14) Seminavis sp2., (13.15) Halamphora yundengensis W.W.Wu, C.P.Chen & Y.H.Gao, (13.16) Halamphora sp1., (13.17,13.18) Halamphora sp nov.
Figure A13. (13.1) Tetramphora decussata (Grunow) Stepanek & Kociolek, (13.2) Amphora immarginata Nagumo, (13.3) Halamphora hyalina (Kützing) Rimet & R.Jahn, (13.4) A. pseudohylina Simonsen, (13.5,13.6) A. helenensis Giffen, (13.7,13.8) A. wisei (M.M.Salah) Simonsen, (13.9) A. marina W.Smith, (13.10). Amphora sp1., (13.11,13.12) Seminavis cf. robusta D.B.Danielidis & D.G.Mann, (13.13) Seminavis sp1., (13.14) Seminavis sp2., (13.15) Halamphora yundengensis W.W.Wu, C.P.Chen & Y.H.Gao, (13.16) Halamphora sp1., (13.17,13.18) Halamphora sp nov.
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Figure A14. (14.1) Halamphora hylina, (14.2) A. pseudhylina, (14.3) A. helenensis, (14.4) Amphora sp., (14.5) H. luciae (Cholnoky) Levkov, (14.6,14.7) Halamphora yundengensis.
Figure A14. (14.1) Halamphora hylina, (14.2) A. pseudhylina, (14.3) A. helenensis, (14.4) Amphora sp., (14.5) H. luciae (Cholnoky) Levkov, (14.6,14.7) Halamphora yundengensis.
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Figure A15. (15.1) Lyrella amphoroides D.G.Mann, (15.2) L. spectabilis (W.Gregory) D.G.Mann, (15.3) Fallacia sp1., (15.4) Fallacia forcipata (Greville) Stickle & D.G.Mann, (15.5) Diploneis crabro Ehrenberg, (15.6) Diploneis sp1., (15.7) D. nitescens (W.Gregory) Cleve, (15.8) Diploneis sp2., (15.9) Diploneis sp3., (15.10) D. papula (A.W.F.Schmidt) Cleve.
Figure A15. (15.1) Lyrella amphoroides D.G.Mann, (15.2) L. spectabilis (W.Gregory) D.G.Mann, (15.3) Fallacia sp1., (15.4) Fallacia forcipata (Greville) Stickle & D.G.Mann, (15.5) Diploneis crabro Ehrenberg, (15.6) Diploneis sp1., (15.7) D. nitescens (W.Gregory) Cleve, (15.8) Diploneis sp2., (15.9) Diploneis sp3., (15.10) D. papula (A.W.F.Schmidt) Cleve.
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Figure A16. (16.1) Diploneis vacillans (A.W.F.Schmidt) Cleve, (16.2) Diploneis sp., (16.3) Falacia Stickle & D.G.Mann sp.
Figure A16. (16.1) Diploneis vacillans (A.W.F.Schmidt) Cleve, (16.2) Diploneis sp., (16.3) Falacia Stickle & D.G.Mann sp.
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Figure A17. (17.1) Mastogloia horvathiana Grunow, (17.2) M. fimbriata (T.Brightwell) Grunow, (17.3) M. angulata F.W.Lewis, (17.4) M. binotata (Grunow) Cleve, (17.5) M. crucicula (Grunow) Cleve, (17.6) M. crucicula var. alternans Zanon, (17.7) M. biocellata (Grunow) G.Novarino & A.R.Muftah, (17.8) M. pumila (Grunow) Cleve.
Figure A17. (17.1) Mastogloia horvathiana Grunow, (17.2) M. fimbriata (T.Brightwell) Grunow, (17.3) M. angulata F.W.Lewis, (17.4) M. binotata (Grunow) Cleve, (17.5) M. crucicula (Grunow) Cleve, (17.6) M. crucicula var. alternans Zanon, (17.7) M. biocellata (Grunow) G.Novarino & A.R.Muftah, (17.8) M. pumila (Grunow) Cleve.
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Figure A18. (18.1) Mastogloia emarginata Hustedt, (18.2) M. ovata Grunow, (18.3) M. ovalis A.W.F.Schmidt, (18.4) M. vasta Hustedt, (18.5) M. adriatica Voigt, (18.6) M. corsicana (Grunow) H.Peragallo & M.Peragallo, (18.7) M. decipiens Hustedt, (18.8) M. cyclops Voigt, (18.9) M. ignorata Hustedt, (18.10) M. asperula Grunow, (18.11) M. lanceolate Thwaites, (18.12) M. laterostrata Hustedt.
Figure A18. (18.1) Mastogloia emarginata Hustedt, (18.2) M. ovata Grunow, (18.3) M. ovalis A.W.F.Schmidt, (18.4) M. vasta Hustedt, (18.5) M. adriatica Voigt, (18.6) M. corsicana (Grunow) H.Peragallo & M.Peragallo, (18.7) M. decipiens Hustedt, (18.8) M. cyclops Voigt, (18.9) M. ignorata Hustedt, (18.10) M. asperula Grunow, (18.11) M. lanceolate Thwaites, (18.12) M. laterostrata Hustedt.
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Figure A19. (19.1) Mastogloia sp1., (19.2) M. paradoxa Grunow, (19.3) M. erythraea Grunow, (19.4) M. regula Hustedt, (19.5) Mastogloia sp2., (19.6) M. cuneata (F.Meister) Simonsen, SEM: (19.7) M. binotata, (19.8) M. corsicana, (19.9) M. crucicula var. alternans, (19.10) M. cuneata.
Figure A19. (19.1) Mastogloia sp1., (19.2) M. paradoxa Grunow, (19.3) M. erythraea Grunow, (19.4) M. regula Hustedt, (19.5) Mastogloia sp2., (19.6) M. cuneata (F.Meister) Simonsen, SEM: (19.7) M. binotata, (19.8) M. corsicana, (19.9) M. crucicula var. alternans, (19.10) M. cuneata.
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Figure A20. (20.1) Pinnularia Ehrenberg sp., (20.2) Caloneis Cleve sp1., (20.3) Caloneis sp2, (20.4) Trachyneis aspera (Ehrenberg) Cleve, (20.5) Plagiotropis cf. lepidoptera (W.Gregory) Kuntze, (20.6) Gyrosigma Hassall sp., (20.7) Haslea Simonsen sp1., (20.8) Haslea sp2., (20.9) Haslea sp3.
Figure A20. (20.1) Pinnularia Ehrenberg sp., (20.2) Caloneis Cleve sp1., (20.3) Caloneis sp2, (20.4) Trachyneis aspera (Ehrenberg) Cleve, (20.5) Plagiotropis cf. lepidoptera (W.Gregory) Kuntze, (20.6) Gyrosigma Hassall sp., (20.7) Haslea Simonsen sp1., (20.8) Haslea sp2., (20.9) Haslea sp3.
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Figure A21. (21.1,21.2) Navicula normalis Hustedt, (21.3) N. ramosissima (C.Agardh) Cleve, (21.4) Navicula sp1., (21.5,21.6) Navicula sp2., (21.7) Navicula sp3., (21.8) Brachysira estoniarum Witkowski, Lange-Bertalot & Metzeltin, (21.9) Proschkinia complanatula (Hustedt) D.G.Mann, (21.10) Climaconeis Gunow sp.
Figure A21. (21.1,21.2) Navicula normalis Hustedt, (21.3) N. ramosissima (C.Agardh) Cleve, (21.4) Navicula sp1., (21.5,21.6) Navicula sp2., (21.7) Navicula sp3., (21.8) Brachysira estoniarum Witkowski, Lange-Bertalot & Metzeltin, (21.9) Proschkinia complanatula (Hustedt) D.G.Mann, (21.10) Climaconeis Gunow sp.
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Figure A22. (22.1) Navivula ramosissima, (22.2) Navicula sp1., (22.3) Navicula sp2., (22.4) N. pavillardii Hustedt, (22.5) Proschkinia complanatula, (22.6) Haslea sp., (22.7) Brachysira estoniarum.
Figure A22. (22.1) Navivula ramosissima, (22.2) Navicula sp1., (22.3) Navicula sp2., (22.4) N. pavillardii Hustedt, (22.5) Proschkinia complanatula, (22.6) Haslea sp., (22.7) Brachysira estoniarum.
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Figure A23. (23.1) Psammodictyon constrictum (W.Gregory) D.G.Mann, (23.2) Nitzschia sp1., (23.3) N. aurariae Cholnoky, (23.4) Nitzschia sp2., (23.5) N. frustulum (Kützing) Grunow, (23.6) Nitzschia sp3., (23.7) Nitzschia sp4., (23.8) Nitzschia sp5., (23.9) N. dissipata (Kützing) Rabenhorst, (23.10) Nitzschia sp6., (23.11) Nitzschia vidovichii Grunow, (23.12) Nitzschia rectilonga Takano.
Figure A23. (23.1) Psammodictyon constrictum (W.Gregory) D.G.Mann, (23.2) Nitzschia sp1., (23.3) N. aurariae Cholnoky, (23.4) Nitzschia sp2., (23.5) N. frustulum (Kützing) Grunow, (23.6) Nitzschia sp3., (23.7) Nitzschia sp4., (23.8) Nitzschia sp5., (23.9) N. dissipata (Kützing) Rabenhorst, (23.10) Nitzschia sp6., (23.11) Nitzschia vidovichii Grunow, (23.12) Nitzschia rectilonga Takano.
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Figure A24. (24.1) Nitzschia amabilis H.Suzuki, (24.2) N. aurarie (24.3) N. frustulum, (24.4) N. inconspicua Grunow, (24.5) Nitzschia sp1., (24.6) Pseudo-nitzschia sp1., (24.7) Pseudo-nitzschia sp2., (24.8) Psammodictyon D.G.Mann sp., (24.9) Nitzschia sp3., (24.10) Nitzschia sp4., (24.11) Nitzschia sp5., (24.12) Nitzschia sp6.
Figure A24. (24.1) Nitzschia amabilis H.Suzuki, (24.2) N. aurarie (24.3) N. frustulum, (24.4) N. inconspicua Grunow, (24.5) Nitzschia sp1., (24.6) Pseudo-nitzschia sp1., (24.7) Pseudo-nitzschia sp2., (24.8) Psammodictyon D.G.Mann sp., (24.9) Nitzschia sp3., (24.10) Nitzschia sp4., (24.11) Nitzschia sp5., (24.12) Nitzschia sp6.
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Phycology 04 00027 g0a25
Figure A25. (25.1) Protokeelia C.W.Reimer & J.J.Lee sp., (25.2) Rhopalodia brebissonii Krammer, (25.3) Rhopalodia sp., (25.4) Auricula complexa (W.Gregory) Cleve, (25.5) Epithemia Kützing sp., (25.6) Surirella Turpin sp1., (25.7) Surirella sp2.
Figure A25. (25.1) Protokeelia C.W.Reimer & J.J.Lee sp., (25.2) Rhopalodia brebissonii Krammer, (25.3) Rhopalodia sp., (25.4) Auricula complexa (W.Gregory) Cleve, (25.5) Epithemia Kützing sp., (25.6) Surirella Turpin sp1., (25.7) Surirella sp2.
Phycology 04 00027 g0a25
Phycology 04 00027 g0a26
Figure A26. (26.1) Rhopalodia brebissonii, (26.2) Auricula Castracane sp., (26.3) Surirella sp.
Figure A26. (26.1) Rhopalodia brebissonii, (26.2) Auricula Castracane sp., (26.3) Surirella sp.
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Phycology 04 00027 g0a27
Figure A27. Dissolved valves (Figure on the left is the regular valve and on the right the valve appears dissolved).
Figure A27. Dissolved valves (Figure on the left is the regular valve and on the right the valve appears dissolved).
Phycology 04 00027 g0a27

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Figure 1. (a) Example of a blue Haslea bloom, H. provincialis, in the open environment (Corsica, France, May 2021), (b) view of living cells of H. provincialis under light microscopy.
Figure 1. (a) Example of a blue Haslea bloom, H. provincialis, in the open environment (Corsica, France, May 2021), (b) view of living cells of H. provincialis under light microscopy.
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Figure 2. Map of the study area including (a) location of the sites in Calvi Bay, (b) location of the sites on the Croatian coast.
Figure 2. Map of the study area including (a) location of the sites in Calvi Bay, (b) location of the sites on the Croatian coast.
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Figure 3. Diatom communities of the 43 samples from the Mediterranean Sea. Samples taken during blue Haslea blooms are highlighted in blue. Controls are blank. The relative abundance of each genus is written in gray. “Others” regroups genera with an RA < 1 %, and the five most abundant genera are written in bold.
Figure 3. Diatom communities of the 43 samples from the Mediterranean Sea. Samples taken during blue Haslea blooms are highlighted in blue. Controls are blank. The relative abundance of each genus is written in gray. “Others” regroups genera with an RA < 1 %, and the five most abundant genera are written in bold.
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Figure 4. PCA on diatom community from 43 Mediterranean samples: (a) clustering of the different sampling sites; (b) the contribution of main genera to the clustering.
Figure 4. PCA on diatom community from 43 Mediterranean samples: (a) clustering of the different sampling sites; (b) the contribution of main genera to the clustering.
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Figure 5. Relative abundance of bacterial communities on turf and Padina sp. on the two sites (O: Oscelluccia and P: port). Blue squares represent samples from bloom conditions. “Others” regroup orders with a relative abundance of less than 1%.
Figure 5. Relative abundance of bacterial communities on turf and Padina sp. on the two sites (O: Oscelluccia and P: port). Blue squares represent samples from bloom conditions. “Others” regroup orders with a relative abundance of less than 1%.
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Figure 6. NMDS of the distribution of bacterial communities in each sample (O: Oscelluccia and S: Port; T: turf; P: Padina sp.). Blue dots represent samples from bloom sites; brown dots are the control samples. A stress value < 0.2 indicates that NMDS can accurately reflect the difference between the samples.
Figure 6. NMDS of the distribution of bacterial communities in each sample (O: Oscelluccia and S: Port; T: turf; P: Padina sp.). Blue dots represent samples from bloom sites; brown dots are the control samples. A stress value < 0.2 indicates that NMDS can accurately reflect the difference between the samples.
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Table 1. Summary table of the different areas and sites sampled. Samples were taken on Padina sp. and turf for further diatom and bacterial analysis.
Table 1. Summary table of the different areas and sites sampled. Samples were taken on Padina sp. and turf for further diatom and bacterial analysis.
AreaYearSiteSubstrate
DiatomDiatom
Calvi Bay2019PortPadina + Turf
MadreporePadina + Turf
2021PortPadina + TurfPadina + Turf
MadreporePadina + Turf
OscelluciaPadina + TurfPadina + Turf
Croatian coast2020ŠunjPadina
DrvenikPadina
Stari GradPadina
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MDPI and ACS Style

Seveno, J.; Witkowski, A.; Car, A.; Gastineau, R.; Sirjacobs, D.; Leignel, V.; Mouget, J.-L. Impact of Blue Haslea spp. Blooms on Benthic Diatom and Bacterial Communities. Phycology 2024, 4, 465-507. https://doi.org/10.3390/phycology4030027

AMA Style

Seveno J, Witkowski A, Car A, Gastineau R, Sirjacobs D, Leignel V, Mouget J-L. Impact of Blue Haslea spp. Blooms on Benthic Diatom and Bacterial Communities. Phycology. 2024; 4(3):465-507. https://doi.org/10.3390/phycology4030027

Chicago/Turabian Style

Seveno, Julie, Andrzej Witkowski, Ana Car, Romain Gastineau, Damien Sirjacobs, Vincent Leignel, and Jean-Luc Mouget. 2024. "Impact of Blue Haslea spp. Blooms on Benthic Diatom and Bacterial Communities" Phycology 4, no. 3: 465-507. https://doi.org/10.3390/phycology4030027

APA Style

Seveno, J., Witkowski, A., Car, A., Gastineau, R., Sirjacobs, D., Leignel, V., & Mouget, J. -L. (2024). Impact of Blue Haslea spp. Blooms on Benthic Diatom and Bacterial Communities. Phycology, 4(3), 465-507. https://doi.org/10.3390/phycology4030027

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