Dead sands: bioerosion of alien foraminiferal shells in a Mediterranean seagrass meadow

Foraminiferans are diverse macroscopic protists abundant in (sub-)tropical seas, often forming characteristic benthic communities known as “living sands”. Numerous species have migrated through the Suez Canal to the Mediterranean, some turning invasive and gradually outcompeting the indigenous species. The most expansive Amphistegina lobifera often creates thick seabed sediments, thus becoming an important environmental engineer. However, little is known about the turnover of its shells in the invaded ecosystems. Using vital staining, stereomicroscopy, scanning electron microscopy, cultivation and 10 DNA fingerprinting, I investigated the vital status, destruction/decomposition and mycobiota of A. lobifera in the rhizosphere of the dominant Mediterranean seagrass Posidonia oceanica in an underwater Maltese meadow (average 284 shells/g, representing 28.5% of dry substrate weight), in comparison with epiphytic specimens and P. oceanica roots. While 78% of the epiphytes were alive, nearly all substrate specimens were dead. On average, 80% of the epiphytes were intact, compared to 21% of the substrate specimens. Abiotic dissolution and mechanical damage played only a minor role, but some bioerosion 15 was detected in 18% and >70% of the epiphytic and substrate specimens, respectively. Few bioerosion traces could be attributed to fungi and the majority probably belonged to photoautotrophs. The seagrass roots displayed fungal colonization typical for this species and yielded 81 identified isolates, while the surface-sterilized substrate specimens surprisingly yielded no cultivable fungi, compared to other 16 identified isolates obtained from the epiphytes. While the epiphytes ́ mycobiota was dominated by ascomycetous generalists also known from terrestrial ecosystems (alongside with, e.g., a relative of the “rock20 eating” extremophiles), the roots were dominated by the seagrass-specific dark septate endophyte Posidoniomyces atricolor and additionally contained a previously unreported lulworthioid mycobiont. In conclusion, at the investigated locality, dead A. lobifera shells seem to be regularly bioeroded by endolithic non-fungal organisms, which may counterbalance their accumulation in the seabed substrate. 25 30 https://doi.org/10.5194/bg-2020-452 Preprint. Discussion started: 8 January 2021 c © Author(s) 2021. CC BY 4.0 License.

interpretation of the history of carbonate depositional environments. In addition, (Kohlmeyer, 1985) suggested that representatives of the genus Amphistegina might be good sources of recent "higher" marine fungi (that colonize and bioerode their shells). Hence, I returned to the same place in May 2018, collected samples of A. lobifera shells from the rhizosphere of the seagrass (and epiphytic specimens + the seagrass roots for comparison) and investigated them using various approaches (vital staining, stereomicroscopy, light and scanning electron microscopy, fungal isolation and DNA fingerprinting): first, to 130 assess the vital status of the A. lobifera specimens as well as their frequency in the substrate and second, to address two central questions of this study, i.e., 1/ what is the fate of dead A. lobifera shells in the P. oceanica rhizosphere and 2/ whether the https://doi.org/10.5194/bg-2020-452 Preprint. Discussion started: 8 January 2021 c Author(s) 2021. CC BY 4.0 License.
fungi inhabiting the seagrass roots colonize the dead shells, thus contributing to their bioerosion. Since the seagrass roots are tightly coupled with a unique spectrum of marine fungi (see above), I hypothesized that these would be the primary bioeroders of dead A. lobifera shells. 135 2 Materials and methods

Sampling
Epiphytic specimens of Amphistegina lobifera, rhizosphere substrate and roots of the seagrass Posidonia oceanica were collected using scuba diving at three different microsites (ca. 10 m apart) at a depth of ca. 6 m at Balluta Bay, St. Julian´s, Malta (GPS: N35.915685, E14.495578) on 28 th May 2018. The epiphytic specimens were collected from P. oceanica leaves 140 and seaweeds growing in the immediate vicinity of the seagrass (Fig. 2c, d) and the substrate containing A. lobifera specimens (volume ca. 50 ml) from the seagrass rhizosphere. All samples were divided in two sub-samples of equal volume, one for (stereo-)microscopic screening and one for mycobiont isolation, and processed as described below.

Screening of Amphistegina shells and Posidonia roots
The sub-samples containing A. lobifera shells were further divided into halves; one half was stained for two weeks with rose 145 Bengal, washed repeatedly with tap water and dried to distinguish alive and dead specimens (Walton, 1952) while the other half was dried and used for counting (to establish the abundance of A. lobifera specimens in 1 g of the dried substrate), weighting (the total weight of A. lobifera specimens in 1 g of the dried substrate), measuring (the diameter of the substrate specimens) and (stereo-)microscopy (to document bioerosion/colonization, dissolution and mechanical damage of the epiphytic + substrate specimens). To measure the diameter of the substrate specimens, random 100 mg of substrate shells per 150 each microsite were separated and the measurements were performed on all shells occurring in three separate fields of view using an Olympus SZX12 stereomicroscope (magnification 12.5×) and the QuickPHOTO MICRO ver. 3.2 software (Promicra, Czechia).
To document bioerosion/colonization, the respective shells were first roughly screened using the stereomicroscope and subsequently, 30 random shells per type and microsite were assessed using a FEI Quanta 200 ESEM scanning electron 155 microscope (FEI Company, USA) in the Low Vacuum mode at room temperature (detailed SEM screening is a lengthy process so the total number of screened shells was primarily limited by the working time available at the SEM microscope). With respect to bioerosion/dissolution, they were sorted out into six qualitative categories, i.e., 1/ intact (=not affected, Fig. 1), 2/ non-bioeroded but partially dissolved, 3/ bioeroded and partially dissolved, 4/ only bioerodedlow level, 5/ only bioerodedintermediate level and 6/ only bioerodedhigh level (of bioerosion). Additionally, surface colonization by macroepiphytes 160 and mechanical damage were recorded (independently of the former six categories) (for illustration see Fig. 3). I did not attempt to determine the respective microborers taxonomically; instead, they were conservatively distinguished into two classes, i.e., fungi and non-fungal organisms. Because the traditional sorting based on the diameter of the borings (e.g., (Perkins and Halsey, https://doi.org/10.5194/bg-2020-452 Preprint. Discussion started: 8 January 2021 c Author(s) 2021. CC BY 4.0 License. 1971)) is not very reliable (see (Golubić et al., 1975)), the borings were assigned to the former class only when intact hyphae were first observed on the shell surface using a stereomicroscope (for illustration see Fig. 4). 165 Random P. oceanica root segments from each microsite were screened for fungal colonization using a compound Olympus BX60 microscope at high magnifications (400× and 1000×) as detailed in (Vohník et al., 2015). In brief, the fine terminal roots were separated from the root system, washed with tap water, their transversal and longitudinal semi-thin sections were prepared using a razor blade and these were mounted in lactoglycerol in glass slides and evaluated for fungal colonization using the compound microscope.

Mycobiont isolation and identification
The protocol for isolation and identification of fungi colonizing A. lobifera shells and P. oceanica terminal roots comprised 175 methods identical to those described in more detail in (Vohník, 2020); this paper also describes their rationale and intuitive troubleshooting. In brief, the low-carbon potato carrot agar (PCA) used for mycobiont isolation was prepared by boiling 40 g of carrots and 40 g of potatoes separately in 500 ml of deionized water for 5 min. The resulting broth was autoclaved at 121°C for 20 min, diluted 1:1 with sterile deionized water, supplemented with agar (10 g/l; HiMedia, India), again autoclaved at 121°C for 20 min and when cooled but still liquid, it was supplemented with Novobiocin sodium salt (50 mg/l; Sigma-Aldrich, 180 Germany) to prevent growth of bacteria. The medium was poured into plastic square 25-compartment Petri dishes and left to solidify under UV light overnight. 50 epiphytic and 50 substrate shells and 50 root segments (ca. 3-4 mm long) were selected randomly from the samples from all three microsites. The shells and the root segments were surface-sterilized 30 s in 10% SAVO (common household bleach; Unilever, Czechia; 100% SAVO contains 47 g kg −1 , i.e., 4.7% sodium hypochlorite = NaClO), 3x washed with sterile 185 deionized water and then transferred onto the surface of the solidified medium in the dishes. Additionally, 25 substrate shells from one microsite were not surface-sterilized but only serially washed with sterile deionized water and then treated as above, serving as a control treatment. The isolations took place during the day of collection. Petri dishes with the shells and root segments were incubated at room temperature in the dark and periodically checked for fungal growth. After six months, all visible fungal cultures were counted, assigned codes and identified as detailed below. As Posidoniomyces atricolor, the 190 dominant root mycobiont of Posidonia oceanica, is notoriously slow-growing (Vohník et al., 2019), the dishes were reexamined after another five months and all new cultures were counted, assigned codes and identified as detailed below.
For mycobiont molecular identification, total DNA was extracted from all fungal cultures producing enough mycelium using an Extract-N-Amp Plant Kit (Sigma-Aldrich, Germany) following manufacturer's instructions. The ITS1-5.8S-ITS2 region (ITS) of the nuclear ribosomal DNA (nrDNA) was amplified using the ITS1F + ITS4 primer pair and the 195 partial large subunit (LSU) nrDNA of some isolates was amplified using the LR0R + LR7 primer pair. The PCR and gel https://doi.org/10.5194/bg-2020-452 Preprint. Discussion started: 8 January 2021 c Author(s) 2021. CC BY 4.0 License. electrophoresis parameters were the same as in (Vohník et al., 2016). The PCR products were purified and sequenced in the Macrogen Europe Laboratory (Macrogen Europe, The Netherlands) using the ITS1, ITS4, LR0R and LR7 primers.
The obtained sequences were screened in Finch TV v1.4.0 (https://digitalworldbiology.com/FinchTV) for possible machine errors and manually edited/trimmed. Where available, the reverse sequences (i.e., those obtained with the ITS4 and 200 LR7 primers) were converted to reverse complement sequences and aligned with the corresponding forward sequences, yielding consensus sequences (contigs) representing the respective fungal isolates. The resulting ITS sequences were subsequently subjected to BLAST searches in GenBank and those not belonging to Posidoniomyces atricolor were aligned in ClustalW (Thompson et al., 1994) implemented in BioEdit v7.2.5 (Hall, 1999). The resulting alignment was used as a matrix ecosystems, e.g., through abiotic dissolution (e.g., (Green et al., 1993)) and bioerosion (e.g., (Cherchi et al., 2012)), transformation of the shells into lime mud, i.e., the important matrix of both recent and ancient calcareous sediments (e.g., 260 (Debenay et al., 1999)), etc. Here, while the abiotic dissolution and mechanical damage contributed only little, the majority (>70%) of the substrate shells showed at least some signs of bioerosion, with 13% being highly bioeroded. This is opposite to, e.g., the findings of (Berkeley et al., 2009) who investigated tropical intertidal sediments in north Queensland, Australia and concluded that the calcareous test degradation during early burial was primarily driven by dissolution, not bioerosion.
However, the reason(-s) for this difference remain unknown. Nevertheless, the data gathered here suggest that bioerosion may, 265 at least to a certain degree, counterbalance the accumulation of alien foram shells in the seabed and thus alleviate the negative impact of the alien foram environmental engineering ((Zenetos et al., 2008); (Yokeş and Meriç, 2009)).
Surprisingly, a great majority of the bioerosion traces seemed to belong to non-fungal organisms (probably cyanobacteria and/or microscopic algae). Congruently, and in contrast to the main hypotheses, not only the substrate shells did not share any fungi with the Posidonia oceanica roots, they did not yield any cultivable fungi at all. This is an unexpected 270 result, because cultivable fungi are ubiquitous in marine ecosystems and regularly colonize calcareous substrates including foram shells (cf. (Kohlmeyer, 1969(Kohlmeyer, , 1984(Kohlmeyer, , 1985). In addition, the epiphytic shells were colonized by fungal ubiquitous generalists as well as specialists and the seagrass roots were regularly colonized by specific symbiotic fungi, including a member of the Lulworthiales that comprise common marine ascomycetes, some of them colonizing foram shells (see (Kohlmeyer et al., 2000)). Nevertheless, a few substrate shells did display apparent signs of fungal colonization by dark septate 275 hyphae (Fig. 4) that actually resembled the mycelium of the dominant P. oceanica root mycobiont (see below). However, an attempt to clone fungal DNA from such shells ended with inconclusive results (data not shown).
The disappearance of cultivable fungi from the substrate shells observed in this study is difficult to explain and one can only speculate about its reasons. For example, since most of the substrate specimens were dead, the respective shells were presumably empty, i.e., without sufficient organic matter to support the fungal growth. However, many marine ascomycetes 280 are notoriously slow-growing (i.e., they need little nutrients), including the dominant P. oceanica root mycobiont (see (Vohník et al., 2019) and references therein) and, e.g., all the foram-associated tropical marine fungi reported by (Kohlmeyer, 1984(Kohlmeyer, , 1985 probably developed on and/or within dead shells. A more likely explanation is allelopathy, a phenomenon common also among marine microorganisms (see (Hellio et al., 2000); (Gross, 2003); (Cepas et al., 2019) and many others). Here, the antagonists could be the (presumably autotrophic) microbioeroders abundant in the substrate shells and/or the fungi inhabiting 285 P. oceanica roots. Indeed, while numerous cultivable fungi have been recently obtained from nearly all P. oceanica tissues, they were absent in the apical parts of the leaves that, however, commonly displayed colonization by microscopic algae/cyanobacteria (B. Soperová and M. Vohník, unpublished results). In addition, while it is still unknown, e.g., how far can reach the mycelium of P. oceanica root-symbiotic fungi, it is interesting to note that their diversity is, at least in the NW Mediterranean Sea, extremely low and dominated by a single mycobiont (Vohník et al., 2016(Vohník et al., , 2017(Vohník et al., , 2019. While data from 290 other seagrasses are too few to allow any robust comparisons, such dominance is extremely rare both in freshwater aquatic https://doi.org/10.5194/bg-2020-452 Preprint. Discussion started: 8 January 2021 c Author(s) 2021. CC BY 4.0 License. and terrestrial ecosystems (e.g., (Vandenkoornhuyse et al., 2002)) and may suggest some kind of antagonism between the dominant root mycobiont and other marine fungi.
Also in this study, the seagrass roots were dominated by P. atricolor, a pleosporalean fungus not known from any other hosts or environments, and the microscopic observations presented here (Fig. 5c, d) provide further indirect evidence 295 that this mycobiont is responsible for the root colonization pattern ubiquitous in the NW Mediterranean Sea (Fig. 5a, see (Vohník et al., 2015)). The seagrass roots additionally yielded a hitherto unknown lulworthioid mycobiont and the epiphytic shells an isolate with affinities to the genus Knufia that comprises highly destructive extremotolerant lithobionts that, e.g., often bioerode Mediterranean historical monuments exposed to outdoor conditions (see (Isola et al., 2016) and references therein). While these isolates represent interesting and potentially important mycobionts and illustrate how little we know 300 about the diversity of marine fungi (see (Gareth Jones, 2011) and references therein), their more detailed taxonomic assignment remained outside the scope and dimensions of this study.

Conclusions
In the first study focused on the fate of A. lobifera during early burial in an invaded ecosystem, I found out that practically all its substrate specimens were dead and regularly bioeroded by presumably photoautotrophic microborers, not marine fungi. 305 Their taxonomic affinities as well as possible antagonistic interactions with the latter remain unknown and beg further investigations. In contrast, the epiphytic A. lobifera specimens yielded a relatively diverse spectrum of mycobionts, at least in comparison with the roots of the seagrass P. oceanica, which comprised both ubiquitous generalist and specialist well-adapted to bioerode calcareous substrates. The switch from fungi in the epiphytic shells to non-fungal organisms in the substrate shells is curious and deserves elucidation, possibly through a study focusing on allelopathic interactions between these two 310 microborer guilds. Nevertheless, a few substrate shells were indeed colonized by unidentified fungus/fungi with dark mycelium and possible future studies on interactions of forams with fungi may consider focusing on foram specimens more intimately associated with seagrass roots.
Author contribution. MV designed the study, organized and performed the sampling on Malta and fungal isolation and identification, analysed the data and wrote the paper.