Fungal parasites infecting N_2-fixing cyanobacteria reshape carbon and N_2 fixation and trophic transfer

Domain: earth_science
Published: 2026-01-02
Editorial outcome: minor_revision

Abstract
Fungal parasites are associated with bloom-forming algae, yet their impact on N2 fixation and the fate of newly fixed nitrogen during cyanobacterial blooms is poorly understood. We report infections on the ecologically important N2-fixing cyanobacterium Dolichospermum (formerly Anabaena) in the Baltic Sea. Using single-cell isotope probing, microscopy, and biogeochemical analyses, we examine how infections affect carbon and N2 fixation and elemental transfer within a natural community. Fungal sporangia infect up to 80% of filaments, mostly targeting storage cells (akinetes, 82% prevalence) and N2-fixing cells (heterocytes, 44%), but rarely vegetative cells (5%). Infections at akinete–heterocyte junctions extract 4- and 10-fold more carbon and nitrogen than those on vegetative cells, reducing host storage by 28% and 56%. Overall, 22% of newly fixed nitrogen is transferred to fungi, comparable to heterotrophic bacteria. Infections also occur in Nodularia and Aphanizomenon, suggesting fungi-like parasitism broadly affects bloom dynamics and the fate of new nitrogen.

Introduction
Algal blooms have significant impacts on aquatic ecosystems and human activities, with both positive and negative consequences. On the one hand, they account for nearly half of global primary production, forming the base of aquatic food webs and supporting fisheries1–3. On the other hand, they can cause oxygen depletion, toxin release, and water quality degradation, threatening ecosystem functioning, biodiversity, fisheries, and recreational activities4,5. Under current climate change, phytoplankton blooms are expanding and intensifying in both coastal and freshwater systems6,7, with harmful effects often outweighing the benefits. For example, in the Baltic Sea, blooms of N2-fixing, partially toxin-producing cyanobacteria have increased in frequency, biomass, and duration since the 1970s8–10. Due to their ability to fix atmospheric N2, diazotrophic cyanobacteria thrive in nitrogen-depleted waters, introducing hundreds of kilotons (180–430 kt) of new nitrogen per year into the central Baltic Sea11,12. This internal nitrogen loading is nearly equivalent to the external nitrogen load from river discharge (480 kt N yr–1)11, exacerbating the vicious cycle that fuels further algal growth, oxygen depletion, and eutrophication13,14. The integration of algal blooms into food webs is typically linked to interactions with zooplankton grazers, heterotrophic bacteria, protists, and viruses15, while cyanobacterial blooms are commonly considered to be regulated by temperature and nutrient availability16,17. In contrast, the role of fungi—particularly parasitic fungi—has largely been overlooked. Mounting evidence, however, suggests that parasitic fungi, belonging mostly to the order Chytridiomycota (hereafter referred to as chytrids), are widespread in coastal18–20 and freshwater ecosystems21,22. While thriving on major phytoplankton groups, including diatoms, cyanobacteria, and dinoflagellates, those fungal parasites infect up to 90% of the phytoplankton host population23–25. Parasitism, such as fungal parasitism, can thereby alter carbon transfer pathways and food web dynamics26,27, whereas its quantitative impact on both N2 fixation and the fate of newly fixed nitrogen during cyanobacterial blooms remains unresolved.

To address this gap, we investigate the exciting interactions between parasitic fungi and N2-fixing cyanobacteria, focusing on infection mechanisms and the quantitative impact on N2 fixation and nitrogen transfer. Using the Baltic Sea as a study site, we apply stable-isotope tracing (13C-bicarbonate and 15N-dinitrogen), single-cell resolution secondary-ion mass spectrometry (SIMS), partial 18S rRNA-gene sequencing, environmental monitoring, microscopy, and biogeochemical analyses. Specifically, we aim to (1) assess the prevalence of fungal infections during summer cyanobacterial blooms, particularly in Dolichospermum (formerly Anabaena); (2) quantify the transfer of carbon and nitrogen from Dolichospermum cells to their fungal sporangia; and (3) evaluate the impact of fungal parasitism on Dolichospermum populations, considering filament integrity and single-cell activity rates.

Results
Dolichospermum long-term observations
The prominent growth season of N2-fixing Dolichospermum spp. in the Southern Baltic Sea extended from June to September between 2012 and 2021 (week 24–40, Fig. 1a). Annual peaks in Dolichospermum biomass predominantly occurred in July (1.9–44.8 µg C L–1, N=6) and, less frequently, in June (18.7 µg C L–1, N=1), August (2.2 and 2.8 µg C L–1, N=2), and September (3.3 µg C L–1, N=1). Between June and September of 2022–2024, Dolichospermum was detected in 33 out of 52 weekly samples. Of these 33 samples, more than half (18 samples) showed signs of infection in the Dolichospermum populations, with the maximum infection prevalence reaching 80% of Dolichospermum filaments (Fig. 1b).

Environmental conditions during isotope-tracer incubations
Environmental conditions at the sampling site on 04 July 2022 reflected typical summer bloom conditions, with low nutrient concentrations (0.23±0.06 µmol PO4 3– L–1, <0.2 µmol NO3 – L–1, <0.05 µmol NO2 – L–1, and 0.65±0.15 µmol NH4 + L–1, N=4 incubation bottles, Supplementary Table S1) and substantial biomass of filamentous N2-fixing cyanobacteria. Specifically, Dolichospermum accounted for 2.0±0.4 µmol C L–1, contributing 34% to the total phytoplankton carbon biomass, while other N2-fixing cyanobacteria contributed less than 1%. Non-N2-fixing phytoplankton species included diatoms (44% of the phytoplankton carbon biomass), dinoflagellates (15%), picocyanobacteria (5%), and other taxa (1%, N=4 incubation bottles, see Supplementary Table S2 for further details).

Characteristics of fungal-infected Dolichospermum
Commonly, Dolichospermum forms chains with differentiated cell types, including vegetative cells (performing oxygenic photosynthesis), heterocytes (performing N2 fixation), and akinetes (resting, energy-reserving cells). To account for this cell differentiation, we considered these three cell types separately during our analyses, as presented in the following.

The Dolichospermum population comprised 88±13 filaments mL–1 or 2298±270 cells mL–1, with the majority of cells being vegetative (91±1%, carbon-fixing cells), followed by heterocytes (7±1%, N2-fixing cells) and akinetes (2±1%, dormant, storage cells). Fungal infections were present in 57±6% of Dolichospermum filaments and 9±3% of their cells (N=12 incubation bottles, Supplementary Table S3). The fungal sporangia were predominantly located at the septal junctions connecting akinetes and heterocytes (Fig. 2a). In total, 82±12% of akinetes and 44±8% of heterocytes were infected, whereas only 5±3% of vegetative cells were infected (N=12 incubations bottles, Fig. 2b). Rhizoids that infected akinetes and heterocytes remained restricted to those cells, whereas those growing into vegetative cells sometimes extended into adjacent vegetative cells. Fungal-infected filaments were, on average, more than twice as long as non-infected filaments (infected: 272±206 µm, N=300; non-infected: 99±70 µm, N=60; Mann–Whitney test, W=138033.5, p=7.68×10–28, rank-biserial correlation=0.43, 95% CI: 0.36–0.49; Fig. 2c). Moreover, akinetes were absent from 95±4% of the shorter, non-infected filaments but from only 36±15% of the infected filaments (N=12 incubation bottles; Mann–Whitney test: W=144, p=3.21×10–5, rank-biserial correlation=1.00, 95% CI: 1.00–1.00). Consequently, the relative abundance of akinetes was significantly lower in non-infected filaments compared to infected filaments (0.5±0.5% vs. 2.1±0.7% of total cells, N=12; Mann–Whitney test: W=5, p=1.21×10–4, rank-biserial correlation = 0.93, 95% CI: 0.83–0.97).

Bacterial colonization of non-infected and fungal-infected Dolichospermum
Non-infected Dolichospermum cells were sparsely colonized by bacteria (0.2±0.6 cell–1, N=100), whereas fungal-infected cells exhibited roughly eight-fold higher colonization (1.7±3.4 cell–1, N=100; ZINB model: count component: z=–4.05, p=5.16×10–5, rate ratio=0.11, 95% CI: 0.04–0.32; Fig. 2d). This increased bacterial colonization was also observed in non-infected cells directly adjacent to fungal-infected cells, where bacterial abundance averaged 0.7±2.4 bacteria cell–1, representing an approximately three-fold increase compared with more distant non-infected cells (N=100). A zero-inflated negative binomial (ZINB) model supported this difference, indicating a significantly higher expected bacterial load in adjacent cells (count component: z=3.90, p=9.68×10–5, rate ratio=9.49, 95% CI: 3.06–29.42). The abundance of free-living bacteria was 1.13±0.14×106 cells mL–1 (N=12 incubation bottles).

Fungal parasite taxonomy and lifecycle
Based on 18S rRNA gene sequencing, the fungal parasite was identified as belonging to the phylum Chytridiomycota. On average, 40±2% of the sequence reads were attributed to Fungi of which 96.4±0.5% were identified as Chytridiomycota (N=4 incubation bottles, Supplementary Fig. S1), which are known to parasitize a broad range of phytoplankton, including, e.g., diatoms, cyanobacteria, and dinoflagellates28–30. The life cycle was typical for early-branching, zoosporic fungi. Zoospores attached to and encysted on the photosynthetic Dolichospermum host. Following encystment, the fungus penetrated the cyanobacterial host and proliferated, conveying its nutrients to the maturing sporangium outside the host. Within this sporangium, new zoospores (ca. 10–20 zoospores) were produced. Eventually, the sporangium opened, releasing the newly generated zoospores, which were capable of swimming and infecting new host cells. The former host cell was left with an empty, collapsed sporangium (Fig. 3a–c).

Single-cell 13C and 15N fixation and transfer
To comprehensively cover the multi-stage life cycle, we analyzed host cells exhibiting the three key stages of sporangial development: (1) zoospore encystment (zoospore loses the flagellum, secretes a cell wall, and adheres to the host cells), (2) mature sporangium, and (3) empty, collapsed sporangium (Fig. 3a–c). Furthermore, we categorized the host cells into distinct types, including vegetative cells, type I heterocytes (adjacent to at least one akinete), type I akinetes (adjacent to one heterocyte), type II heterocytes (enclosed by two vegetative cells), and type II akinetes (enclosed by two vegetative cells) (Fig. 3d). Our single-cell isotope analyses addressed four central questions: Does the fungal parasite acquire more cyanobacterial-derived carbon and nitrogen when infecting septal junctions between type I akinetes and heterocytes, compared to less preferential sites such as vegetative cells, type II akinetes, and type II heterocytes? How efficiently does the fungal parasite siphon carbon and nitrogen from its cyanobacterial host? Is this cell-to-cell transfer comparable to that of bacteria? Are cyanobacterial cells of the same cell type (vegetative, heterocytes, or akinetes) with high carbon and N2 fixation activity preferentially targeted for infection by the fungal parasite? Does fungal infection reduce carbon and N2 fixation rates and storage in Dolichospermum cells?

Site-specific acquisition of cyanobacterial carbon and nitrogen by the fungus
The fungal parasite exhibited the highest 13C and 15N enrichment in mature sporangia infecting type I akinete–heterocyte junctions. Specifically, the 13C and 15N enrichment was 4-fold and 10-fold greater in these sporangia (1.42±0.33 13C and 0.25±0.10 15N atom% excess, N=33) compared to sporangia infecting vegetative cells (0.32±0.33 13C and 0.02±0.01 15N atom% excess, N=41, Supplementary Fig. S2). Interestingly, sporangia infecting type II heterocytes or akinetes—which were not adjacent to each other but enclosed by vegetative cells—showed low enrichment levels, comparable to vegetative cells (type II heterocytes: 0.35±0.54 13C and 0.04±0.05 15N atom% excess, N=8, type II akinetes: 0.19±0.09 13C and 0.02±0.01 15N atom% excess, N=5, Supplementary Fig. S2).

Pairwise comparisons revealed that sporangia at akinete–heterocyte junctions were enriched in both 13C and 15N to levels exceeding those of their host cells (Fig. 4). Specifically, the 13C atom% excess in mature sporangia was 1.2-fold (1.2±0.3, N=24) and 4.0-fold (4.0±2.1, N=28) greater, and the 15N atom% excess was 2.3-fold (2.3±0.5, N=24) and 3.6-fold (3.6±1.5, N=28) greater than in their type I akinete and heterocyte host cells, respectively (Fig. 4a, b). In contrast, sporangia associated with vegetative cells were significantly less enriched in both isotopes than their host cells (paired Wilcoxon signed-rank test: V=1485, p=1.67×10–10, matched rank-biserial correlation=–1, 95% CI: –1 to –1; Fig. 4a, b). Their 13C and 15N enrichment was only 0.2-fold (0.21±0.21) and 0.1-fold (0.14±0.05, N=54) the enrichment found in their vegetative host. Carbon and nitrogen were, therefore, efficiently transferred from type I akinetes and heterocytes to the fungal parasite, even exceeding the newly fixed carbon and nitrogen retained within the host cells (Fig. 4c, d). Conversely, newly fixed carbon and nitrogen were poorly transferred from vegetative cells to parasitizing sporangia.

13C and 15N enrichment levels were similar among bacteria associated with different Dolichospermum host cell types: those associated with akinetes showed 0.67±0.42 13C and 0.075±0.075 15N atom% excess (N=12), with heterocytes, 0.48±0.44 13C and 0.069±0.048 15N (N=9), with sporangia, 0.55±0.39 13C and 0.070±0.045 15N (N=38), and with vegetative cells, 0.62±0.44 13C and 0.095±0.082 15N (N=63). On average, enrichments across all Dolichospermum-associated bacteria were 0.59±0.42 13C atom% excess and 0.08±0.07 15N atom% excess (N=122). This was at least 2.4-fold lower than in mature sporangia infecting type I akinete–heterocyte junctions, but at least 1.7-fold higher than in sporangia infecting vegetative cells (Supplementary Table S4). In comparison, free-living bacteria showed considerably lower enrichment, with 0.18±0.11 13C and 0.043±0.018 15N atom% excess (N=11).

Cells within the same host type (vegetative, heterocytes, or akinetes) showed no significant differences in carbon and N2 fixation-based growth rates between non-infected host cells and cells that had just been infected, as indicated by an encysted zoospore (p>0.05, two-sample Student’s t test and Mann–Whitney, see Supplementary Fig. S3 for details). Thus, there was no preference for infection based on the level of host activity within the same cell type.

Impact of fungal infection on cyanobacterial carbon and N2 fixation
Fungal infection significantly reduced net carbon incorporation (retention) rates by 28% in type I akinetes (from 4.5±1.1 pmol C cell–1 d–1, N=104 to 3.2±1.2 pmol C cell–1 d–1, N=65; two-sided t-test: t(167)=–7.08, p=3.76×10–11, Cohen’s d=–1.12, 95% CI: –1.45 to –0.79) and by 23% in type I heterocytes (from 0.10±0.05 pmol C cell–1 d–1, N=55 to 0.08±0.04 pmol C cell–1 d–1, N=62; two-sided Wilcoxon rank-sum test: W=1232.5, p=0.00994, rank-biserial correlation=–0.28, 95% CI: –0.46 to –0.07). The effect on vegetative cells, however, was insignificant (non-infected: 0.39±0.08 pmol C cell–1 d–1, N=3527 and fungal-infected: 0.38±0.10 pmol C cell–1 d–1, N=67; two-sided Wilcoxon rank-sum test: W=10 746.5, p=0.244, rank-biserial correlation=–0.24, 95% CI: –0.57 to 0.16, Fig. 5a–c). Similarly, net nitrogen incorporation was reduced by 56% in type I akinetes (from 0.71±0.45 pmol N cell–1 d–1, N=104 to 0.31±0.21 pmol N cell–1 d–1, N=65; two-sided Wilcoxon rank-sum test: W=1154, p=6.36×10–13, rank-biserial correlation=–0.66, 95% CI: –0.75 to –0.54) and by 30% in type I heterocytes (from 0.018±0.011 pmol N cell–1 d–1, N=55 to 0.013±0.010 pmol N cell–1 d–1, N=62; two-sided Wilcoxon rank-sum test: W=1030, p=2.24×10–4, rank-biserial correlation=–0.40, 95% CI: –0.56 to –0.21), while no significant change was evident in vegetative cells (non-infected: 0.04±0.02 pmol N cell–1 d–1, N=3527 and fungal-infected: 0.04±0.02 pmol N cell–1 d–1, N=67; two-sided Wilcoxon rank-sum test: W=13073, p=0.720, rank-biserial correlation=–0.07, 95% CI: –0.44 to 0.32). The net C:N incorporation ratio (mol: mol) increased 1.6-fold in infected type I akinetes (from 8.3±4.4, N=104 to 12.8±6.8, N=65; two-sided Wilcoxon rank-sum test: W=4942, p=4.51×10–7, rank-biserial correlation=0.46, 95% CI: 0.31 to 0.59), but remained unchanged in type I heterocytes (non-infected: 6.1±2.3, N=55, fungal-infected: 7.1±3.5, N=62; two-sided Wilcoxon rank-sum test: W=2015.5, p=0.0905, rank-biserial correlation = 0.18, 95% CI: –0.03 to 0.38) and vegetative cells (non-infected: 11.1±6.3, N=3527, fungal-infected: 9.8±3.9, N=67; two-sided Wilcoxon rank-sum test: W=14583.5, p=0.869, rank-biserial correlation=0.03, 95% CI: –0.35 to 0.41, Fig. 5d–f).

Methods
Sampling and stable isotope incubations
We collected water samples during summer (July 4th, 2022, 21:00, UTC+2) at the coast of the Southern Baltic Sea (Belt Sea, Heiligendamm, 54°08'46.7''N, 11°50'36.1''E, 0.6m sampling depth, 5m max. water depth) using a Niskin water sampler (Single PWS 1.7 L, Hydro-Bios, Germany). Water temperature was 18.7°C and salinity 9.5 (measured with CTD 48 SSD A Sea & Sun Technology). The samples were transported to the nearby laboratory, and incubations started two hours after sampling.

We filled the water headspace-free into twelve 1 L Duran® bottles, amended nine of the bottles with pre-dissolved 15N-N2 (15N-N2 98%, 29817-79-6, Eurisotop, Germany) and 13C-DIC (dissolved inorganic carbon, NaH13CO3, 98%, 372382, Sigma-Aldrich, Germany), and closed them with a membrane lid (Duran® silicone septum, PTFE coated). The isotope amendment was 3.9±0.1 13C atom% excess in the DIC pool and 0.67±0.07 15N atom% excess in the N2 pool (N=9, see below Bulk isotope analyses). Subsequent incubations lasted for 10 h in darkness (10 h), and 10 h in darkness followed by 11 hours in light (21 h) inside a growth chamber (KBW 400, BINDER, Germany) at 18°C with 0 µE m–2 during darkness and 40 µE m–2 during light. At each time point, we sampled three replicate bottles with isotope enrichment and one control bottle without enrichment for the following analyses.

Nutrients
Samples of 80 mL were filtered through glass fiber filters (Advantec™ Grade GF75, pre-combusted at 480°C for 5 h), the filtrate was transferred into HCl-washed (pH=4) 30 mL-HDPE vials and frozen at –20°C. Nutrients (PO4 3–, NO3 –, NO2 –, and NH4 +) were measured colorimetrically according to Grasshoff et al55. using a Seal Analytical QuAAtro MT3B automated constant flow analyzer for seawater (lowest standards: 0.005 µM PO4 3–, 0.2 µM NO3 –, 0.05 µM NO2 –, and 0.5 µM NH4 +, precision 5%). Samples for dissolved organic carbon (DOC) and dissolved nitrogen (DN) were stored in 20 mL glass vials at –20°C and analyzed through combustion catalytic oxidation (lowest standards: 25 µmol DOC L–1 and 5 µmol DN L–1).

Phytoplankton abundance and infection prevalence
We preserved 50 mL samples with Lugol’s solution (100 g potassium iodide and 50 g iodine in 1 L, pH=7, 50 µL added per 1 mL sample) and stored them at 4°C. Prior to microscopy, we decolorized Lugol samples with sodium thiosulfate (Na2S2O3, final conc. 7.6 mM, CAS# 7772-98-7, Sigma Aldrich) (Pomroy 1984) and stained with Calcofluor White (5 µg mL-1, CFW, Fluorescent Brightener 28, Sigma Aldrich) and Wheat Germ Agglutinin (5 µg mL–1, WGA, Alexa Fluor™ 488 Conjugate, Thermo Fisher Scientific Inc., MA, USA)56. Samples were inspected in Utermöhl chambers at 200× (Nikon, CFI Super Fluor 20×) and 600× magnification (Nikon, CFI P-Apo 60×) with an inverted epifluorescence microscope (Nikon Eclipse Ti2, Nikon Imaging Inc., Tokyo, Japan) using a long-pass DAPI filter (DAPI-50LP) for CFW signals and a FITC filter for WGA signals. We recorded (i) Dolichospermum cell abundances (cells mL–1), (ii) fungal infection prevalences in Dolichospermum filaments (%), (iii) infection prevalences in the three different Dolichospermum cell types, i.e., vegetative cells, heterocytes, and akinetes, and (iv) Dolichospermum filament lengths (µm, cell chains with less than four cells were not counted as filaments). We additionally analyzed samples from the initial time point for (v) abundances of co-occurring autotrophs and mixotrophs (cells mL–1) and biomass (µmol C L–1)57. Since our samples were derived from the water column, we classified Dolichospermum as the planktonic Dolichospermum (in contrast to the benthic Anabaena)58, in line with the classification used in the Baltic Sea monitoring (Baltic Marine Environment Protection Commission, Helsinki Commission, HELCOM).

Bacterial abundances
Samples for bacterial abundance were fixed with paraformaldehyde (PFA, final conc. 1.5%) and filtered onto 5.0 µm (30 mL) and 0.2 µm filters (3 mL) (Whatman® Nucleopore™ Track-Etched, 25 mm) for separating Dolichospermum-associated and free-living bacteria, respectively. Filters were incubated with CFW (1 µg mL–1) and SYTOX Green (final conc. 0.1 µg mL–1, NucGreen™ Dead 488, DNA-binding) for 10−15 min in darkness. Excess stain was rinsed off during 1 min-washing steps with PBS, MilliQ, 50% EtOH, and 100% EtOH. Filters were air-dried and mounted with a 1:1 mix of Vectashield® (Antifade Mounting Medium, Vector Laboratories, USA) and CitiFluor™ (AF1, Science Services GmbH, Germany). Host-associated bacteria were counted manually on 200 host cells (25 non-infected and 25 infected cells per replicate bottle), as well as their two neighboring cells. Free-living bacterial abundances were counted in 156×156 µm areas (N=25 per replicate bottle) using ImageJ2 (v1.54f)59. Stacked images were converted to 8-bit format, and a threshold was applied (gray value 15−255) for automated particle analysis.

Fungal community composition
To verify the taxonomy of the unknown fungal parasite, we resolved the fungal community using 18S rRNA-gene amplicon sequencing. We filtered 500 mL onto 5.0 µm PC filters (Whatman®, Nucleopore, 25 mm), which were transferred into cryovials, placed into liquid N2, and then frozen at –80°C. To detach cells from the filters, filters were vortexed with 0.5 mm beads and 650 µL CTAB (Cetyl-trimethylammonium bromide) lysing buffer (Tris–HCl 100 mM, pH 8.0, NaCl 1.4 M, EDTA 20 mM, CTAB 2%) was added60. After the addition of 65 µL 10% SDS and 10.6 µL Proteinase K (conc: 2 mg mL–1, 30 units mg–1), tubes were gently shaken and placed into a heating block at 60°C for 1 h. Tubes were thereafter exposed to a bead-beating step (FastPrep-24™, MP Biomedicals, time: 40 s, speed 6.0 ms–1) and then loaded with Phenol−Chloroform−Isoamyl alcohol (P:C:I, 25:24:1). We vortexed the mixture for 5×3 sec, mixed by hand to ensure phenol had maximum contact with proteins, and centrifuged for 10 min at 10,000×g and 4°C. The upper phase was transferred into a new 1.5 mL tube, and the P:C:I extraction was repeated. A final cleaning was done using chloroform:isoamyl alcohol (24:1) before the samples were incubated in isopropanol at –20°C overnight. Tubes were centrifuged at 16,000×g and 4°C for 30 min. The supernatant was removed, and the pellet was washed with ice-cold 70% ethanol. Steps of centrifugation at 17,000 xg for 10 min, supernatant removal, and EtOH washing were repeated twice. The remaining EtOH was removed, and the dried pellet was reconstituted in 30 µL PCR water at 37°C at 450 rpm for 30 min. Prior to DNA sampling and extraction, we cleaned all disposables and surfaces with 1.2% bleach and 96% EtOH, and additionally heat-cleaned (480°C, 5 h) the glassware to minimize DNA contamination.

The 18S rRNA-encoding DNA was amplified using the primer set FF390/FR-1 (nu-SSU-1333/nu-SSU-1647, V7/8 region)61,62. To reduce the co-amplification of Stramenopiles, an annealing-blocking oligonucleotide was used62. DNA was amplified using a Master Mix (Premix Ex Taq™ DNA Polymerase Hot Start Version, TaKaRa, Japan). Each PCR program included 240 s at 94°C, 30 cycles of 30 s at 94°C, 60 s at 44.5°C, and 90 s at 72°C, and finally 300 s at 72°C. As a positive control, we used a microbial community DNA standard (D6306, Zymo Research Europe) to reveal possible biases throughout the workflow and analysis. Duplicate samples (i.e., technical replicates) were created by adding the same DNA extract for amplification. Amplified PCR products were purified using AGENCOURT® AMPURE® XP beads (A63881, Beckman Coulter, CA, USA) and a magnetic separator. Illumina Sequencing (MiSeq, v3 600 cycles kit, 2×300 bp) was performed at the Deep Sequencing Facility of the DRESDEN-concept Genome Center (Dresden, Germany).

Primers were removed from the sequences with cutadapt (v2.8). Truncation, quality filtering, dereplication, pooling, and chimera removal were done with dada2 (1.34.0)63 in RStudio (2024.12.0)64. For quality filtering, a maximum of two erroneous bases was allowed for forward and reverse reads, and ten base pairs were used as an overlap when merging forward and reverse reads. For taxonomic assignment of each amplicon sequence variant (ASV), we used the PR2 SSU database (pr2_version_5.1.0_SSU_dada2) as a reference. For comparison, we also used the SILVA SSU database (silva_SSUfungi_nr99_v138_2_toGenus_trainset). Both databases classified 39% of the ASVs as Chytridiomycota. Moreover, SILVA assigned 99% of those reads to class-level Chytridiomycetes Incertae Sedis, while PR2 classified 3% as Rhizophydiales, 0.04% as Chytridiales, 0.04% as Lobulomycetales, and 0.02% as Polychytriales, while the remaining reads remained unclassified at the class/order level (default minBoot=50). Additional fungal reads were assigned to Ascomycota, Basidiomycota, and Zoopagomycota (1.7%, 0.2%, and 0.9% of fungal reads). Likely contaminants were detected and removed using decontam (1.26.0)65. Relative abundances of ASV counts were plotted after normalizing for sample depth with DESeq2 (1.46.0)66.

Bulk isotope analyses
We filtered 700 mL onto pre-combusted filters (Advantec™ Grade GF75 Glass Fiber Filters, pre-combustion at 480°C for 5 h) and stored them at –20°C. Filters were later dried at 50°C, fumed over HCl, pelletized into tin cups, and analyzed for 13C particulate organic carbon (POC) and 15N particulate organic nitrogen (PON) using an elemental analyzer IRMS (EA-IRMS, precision ±0.2‰ for 13C and ±0.3‰ for 15N) at the Stable Isotope Facility (SIF), UC Davis (California, US). The filtrate was transferred headspace-free into 12 mL glass Exetainer® vials (#739W, Labco Limited, Wales, UK), preserved with zinc chloride (ZnCl2, final concentration 0.05% w/v, CAS 7646-85-7, Merck), and stored at room temperature for the determination of 13C-label% in the DIC pool and 15N-label% in the dissolved N2 pool. The 13C-label% of DIC was quantified by trace-gas IRMS after converting DIC to CO2 using 85% phosphoric acid at the Stable Isotope Facility (SIF, UC Davis). The 15N-label% of dissolved N2 was analyzed using a gas concentration and purification system interfaced with an isotope-ratio mass spectrometer in the same laboratory (raw data can be found in Supplementary Dataset 1). Additional technical details on mass spectrometer analyses at UC Davis are provided in Supplementary Note 3. Each analysis included four replicate vials (three with isotope amendment and one control without amendment). Carbon and N2 fixation or incorporation rates were calculated as follows (example shown for nitrogen):

N fixation rate = (APN - APN0) / (AN - APN) * [PN] / Δt (1)

where APN is the fractional 15N-enrichment in the particulate nitrogen (PN) pool following the incubation (tn), APN0 the fractional 15N-enrichment in the particulate nitrogen (PN) pool before the incubation (t0), AN the fractional 15N-enrichment (atom%) of the nitrogen source pool, PN the concentration of PN at the end of the incubation period, and Δt the incubation duration67.

Single-cell isotope analyses
Samples (30 mL) were fixed with PFA (final conc. 1.5%), collected onto 5.0 µm filters (Isopore, TMTP02500, Merck, Darmstadt, Germany), and stored at –20°C. Before analyses, filters were cut into circa 4×4 mm sections, glued onto adhesive carbon tape on a 1-inch glass slide, and coated with a 30 nm gold layer (Cressington 108 Auto Sputter Coater, Watford, UK). Cells were imaged at 800× and 1200× magnification under a scanning electron microscope (SEM, Merlin VP compact, Zeiss, Germany) to identify and later relocate non-infected and infected Dolichospermum cells during SIMS analyses. Given a higher sample throughput, Dolichospermum cells and associated sporangia were analyzed using a large-geometry secondary ion mass spectrometer (IMS 1280, CAMECA, Gennevilliers, France) at the NordSIMS facility (Natural History Museum, Stockholm, Sweden) following26. Areas of interest (80×80 µm) were pre-sputtered with a primary cesium ion beam (Cs+, 5 nA) for 3.5 min to remove the gold coating and cell walls. Data acquisition was done on 70×70 µm rasters (256×256 px resolution) with a 50 pA Cs+ beam (spatial resolution 500–1000 nm), utilizing the synchronized secondary ion beam raster (dynamic transfer optical system, DTOS) to maintain high transmission and high mass resolution and to facilitate reconstruction of the ion image by the instrument software. The secondary ion signals of 12C14N-, 12C15N-, and 13C14N- were recorded by a low-noise (<0.005 counts per second) ion counting electron multiplier at least 50 planes using a peak-switching routine with dwell times of 15, 76, and 31 µs px–1 plane–1, respectively. The mass resolution was 12,000 (M/ΔM).

The analysis of bacteria (Dolichospermum-associated and free-living bacteria) required a higher spatial resolution and was thus done on a NanoSIMS 50L (50−100 nm resolution, CAMECA, Gennevilliers, France). Areas of interest (30×30 µm) were pre-sputtered with a primary Cs+ ion beam (600 pA) for 1.5 min. Data acquisition was done on 25×25 µm rasters (512×512 px resolution) with a 1 pA Cs+ beam, utilizing the synchronized secondary ion beam raster (continuous, magnetic sector/DTOS). The secondary ion signals of 12C13C-, 12C12C-, 15N12C-, 14N12C-, and 31P- were recorded by a low-noise (0.541 counts per second) ion counting electron current pre-amplifier over 60 planes in multi-collection mode with a dwell time of 250 µs px–1 plane–1. The resolution of each mass was adjusted toward the best peak identification before each measurement. The mass resolution was 10,000 M/ΔM for 12C12C-, 15N12C-, and 14N12C- (given by CAMECA software).

Regions of interest (ROI) were selected from previously taken SEM images and drawn manually on the 12C14N ion images using the software WinImage (v4.8). Free-living bacteria were defined as those located at least 10 µm distant to Dolichospermum cells (equal to approximately twice the Dolichospermum diameter). Isotope ratios for each ROI were calculated based on the cumulative ion counts (cts) of all planes as 13C/12C= cts 13C14N/ cts 12C14N- (IMS 1280), 13C/12C= cts 12C13C-/cts 12C ×0.5 (NanoSIMS), and 15N/14N= cts 15N12C-/cts 14N12C- (IMS 1280 and NanoSIMS). Isotope ratios with a standard error exceeding twice their Poisson statistics error (σ ratio), and with a relative standard deviation above 25% across all planes, were rejected. Ratios were corrected for instrument mass fractionation using our EA-IRMS data obtained from GF75 control filters (see above), following68 and converted to atom% excess (APE), i.e., excess label% in a given cell type relative to non-enriched control cells. We used isotope atom% excess values to calculate C- and N-specific assimilation rates (d–1) of host cells, and multiplied those by Dolichospermum cell-specific C- and N-contents (pmol cell–1) (Supplementary Table S6), respectively, to obtain cell-specific rates (pmol cell–1 d–1), C- and N-specific rates (d–1)69,70 (see Supplementary Note 4 for equations). We refer to these rates (pmol cell–1 d–1) as net carbon or nitrogen incorporation rates, as they represent the amount of newly fixed carbon and nitrogen retained in host cells after some was siphoned off by the parasite or lost through other pathways during the incubation. Preparing samples for SIMS analyses is known to cause isotope dilution71,72, which we did not correct for due to an uncertain dilution factor. Our sample preparation steps, however, were kept minor with only PFA preservation, known to cause moderate isotope dilution of approximately 10%71. Herein reported ratios should thus be considered conservative estimates.

Dolichospermum long-term observations
To provide an overview of the long-term Dolichospermum biomass, we used long-term monitoring data obtained from the same sampling site from 1998 to 2021 (ODIN database, https://odin2.io-warnemuende.de/). Fungal infections are not included in this monitoring program. We thus collected additional samples weekly at the Heiligendamm site during 2022−2024 (supported by the IOW monitoring program), to monitor fungal infections in Dolichospermum populations. Depth-integrated samples were collected from the upper water column (0–1 m) with a vertical plankton net (10 µm mesh size, Apstein 50, Hydro-Bios GmbH, Germany), preserved with Lugol’s solution (pH=7) and stored in 100 mL brown glass bottles at 4°C. Using microscopy, we counted the relative abundance of fungal-infected and non-infected filaments, inspecting 100 Dolichospermum filaments (if present) as described above (see Phytoplankton abundance and infection prevalence).

Statistical analyses and software
Statistical differences between two samples were calculated using the Mann–Whitney test for non-normally distributed data, the Welsh test for normally distributed data with non-equal variance, and the t-test for normally distributed data with equal variance. Normal distribution was verified using the Shapiro test and data variance with the F-test. Statistical differences between multiple groups were determined with Tukey’s HSD test (for normally distributed data populations) and the Kruskal–Wallis test (if normal distribution was rejected, with Bonferroni correction for p-value adjustment). All statistical tests were two-sided. Count data of associated bacteria were dominated by zeros and thus modeled using a zero-inflated Poisson (ZIP) and Zero-Inflated Negative Binomial (ZINB) model. The model with the lower Akaike Information Criterion (AIC) was given preference. Tests were run in RStudio 2024.12.0 using the packages tidyverse73, agricolae74, pscl75,76, and MASS77. Figures were created in RStudio 2024.12.0 (R version 4.4.2), Origin 2022, and CorelDRAW 2020. Microscope images were taken with the Nikon NIS-Elements (v. 5.21).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript of Feuring et al describes the discovery of cyanobacteria infecting fungal parasites and the trophic transfer of C and N to the fungi by the N2-fixing cyanobacteria. To my knowledge, this is the first study to demonstrate this association and quantify its elemental cycling impact in nature. The authors observed close associations between the fungal parasites and the filamentous cyanobacteria over several years in a coastal area prone to algal blooms. They demonstrate with very nice stable isotope labeling experiments that the fungi take up N and C directly from the cyanobacteria, and have a stronger preference for attacking the heterocyst and akinetes (compared to vegetative cells). It is a great study, novel results, that are important for the fields of microbial ecology and biogeochemistry in general and should be of broad interest to the readership of Nature Communications. I only have one suggestion for an additional analysis - the authors should make a 18S rRNA gene phylogenetic tree (because they have the sequence) from their parasite that they discovered. This will show how it is related to other parasites that have already been discovered. Is it a new species? Or one that is already known? The tree will answer this important question. I have just a few other minor comments and suggestions, hopefully this helps the authors when revising the manuscript:\nComments:\nlines 165-176: Since the whole 18S rRNA gene sequence has been obtained, the authors should provide a more precise taxonomic analysis. I recommend they do a phylogenetic analysis to see where these particular parasites fall within the Chytridiiomycota tree, and whether it is a novel group or a group that is already known from other locations/studies.\nstarting on line 200: It is a bit unconventional to write a scientific results and discussion section with this Q1 , Q2, et c formatting. I suggest making these individual paragraphs with section headers.\nlines 212-234: The discovery of the fungal preference for the N2-fixing heterocyst and N-rich akinete storage cells over the vegetative cells is extremely interesting! A remarkable result indeed. This high nitrogen assimilation by the marine fungi associated with the N2-fixing heterocysts makes me wonder, if organic nitrogen assimilation is an important part of the ecology of marine fungi in general. It reminds me of similarly high organic N assimilation activity inferred for marine fungi in the Namibian upwelling zone (doi.org/10.1038/s41396-021-01169-5). Could the authors comment on organic N assimilation more generally as relates to marine fungal ecology in the discussion? And, also - what the implications of fungal organic N assimilations might be for the marine N cycle?\nlines 309-311: This is a really interesting discovery\nlines 328-329: This is where the phylogenetic tree of the 18S gene from the chytrids studied her would be useful, R crass and R anabaena should also be represented in the tree. Is it the same species or different?\nlines 331-332: This can be easily resolved using a simple 18S phylogenetic tree analysis.", "reviewer_2": "Feuring and colleagues present their study showing that fungal parasites significantly impact nitrogen cycling during cyanobacterial blooms by infecting key cell types in Dolichospermum, extracting substantial amounts of host-derived carbon and nitrogen. These infections divert newly fixed nitrogen to fungi, rivaling bacterial uptake, and suggest fungal parasitism is a widespread and influential factor in bloom dynamics in the Baltic Sea.\nThough prevalent in aquatic ecosystems, we still only have a limited understanding of chytrid ecology and studies like this are therefore critical. The team are experts in applying sophisticated isotope techniques to study chytrid-algal interaction and this approach is used well here.\nA generally well-written manuscript. I have a few points to consider.\nTo what extent the novelty of the system needs stating more clearly. This a new marine observation but has it been shown to some extent already in freshwater ecosystems?\nTo what extent is this a local phenomenon for the Baltic Sea? Could a statement be made of the wider significance of this work?\nThe early results on Dolichospermum distribution – is this not already known? This feels rudimentary.\nThe selection of specific cell types is fascinating and a key result. Could you observe the rhizoids inside the cells? Do they stay in the targeted cell-type or move to the next?\nEncysted zoospore feels contradictory. Can you define? E.g. flagellum retracted?\nThe observation of more bacteria associated with infected cells. Brilliant result. Could more be explained on this?\nI am not familiar with isotopic techniques used, so cannot comment on the validity of the approach."}, "author_response": "We thank the reviewer for the careful evaluation of our manuscript, the positive feedback, and constructive suggestions, which have helped us to improve the quality of the work.\nBelow, we provide a detailed, point-by-point response to each comment.\nLine numbers correspond to those in the final document without track changes. Additional edits were made to ensure compliance with the Nature Communications formatting guidelines (a title without punctuation, shortened abstract, etc.).\nComments:\nlines 165-176: Since the whole 18S rRNA gene sequence has been obtained, the authors should provide a more precise taxonomic analysis. I recommend they do a phylogenetic analysis to see where these particular parasites fall within the Chytridiiomycota tree, and whether it is a novel group or a group that is already known from other locations/studies.\nResponse: We thank the reviewer for this valuable suggestion. Our study did not target the full 18S rRNA gene, but specifically the V7/8 region amplified with primers FF390/FR-1 (nu-SSU-1333/nu-SSU-1647; see line 467 f.). These primers were chosen because they are well established for fungal community profiling (Banos et al 2018, Vainio and Hantula 2000), compatible with short-read sequencing, and allow the use of blocking oligonucleotides to reduce non-fungal amplification. The design of our study was therefore aimed at providing an overview of the fungal community composition in our environmental samples rather than in-depth phylogenetic resolution.\nTo nevertheless address the reviewer’s point, we explored the phylogenetic placement of the most abundant fungal ASV (ASV_8, representing 87% of Chytridiomycota-assigned reads).\nA BLAST search on NCBI revealed a best hit with 99.7% identity to an uncultured eukaryote clone (HQ873384.1) (Prenafeta-Boldu et al 2014). Interestingly, this highly similar sequence was derived from a coastal dune system, but neither parasitic fungi nor phytoplankton/cyanobacteria were mentioned in the study (Prenafeta-Boldu et al 2014). Additional hits (89–91% identity) matched various uncultured Chytridiomycota and Rhizophydiales.\nWe further constructed trees directly on NCBI using (i) the 100 best BLAST matches and (ii) sequences of chytrids known to infect phytoplankton (Van den Wyngaert et al 2022) (Fig. 1 below). In the latter case (ii), ASV_8 clustered broadly within chytrid-related fungi, but with relatively low percent identities (84–92%) suggesting resolution at the order or family level, rather than genus or species (Table 1, Fig. 1B below). We also included a chytrid sequence derived from infected Dolichospermum akinetes in a lake, similar to our sample, but this sequence showed only 90.7% identity with ASV 8 (OL869110.1 in Table 1), and 75.6–90.7% identity with all ASVs. We added this information to line 273 ff.\nTo our knowledge, the V7/8 region does not provide sufficient resolution for robust phylogenetic inference. Moreover, a publishable tree would require curated reference alignments beyond the objectives of this study. We thus prefer not to include a detailed phylogeny in the manuscript. However, our additional analyses, together with published data, highlight that the taxonomic term “chytrid” may be too narrow for our dataset. In the revised version, we therefore use more cautious terminology such as “fungal parasites,” “zoosporic fungi,” or “chytrid-like fungi.”\nTable 1. Percent identity of ASV_8 with selected publicly available sequences of chytrids known to parasitize various phytoplankton hosts (data obtained from Van den Wyngaert et al 2022).\nGenBank Sequenced Organism Identified Phytoplankton Host Percent Identity\naccessionno.\nOL869111 uncultured Zygophlyctidales Asterionella 91.6\nOL869110 uncultured fungus Dolichospermum (akinete) 90.9\nOL869119 uncultured Staurastromyces sp. Staurastrum 90.9\nOL869115 uncultured Rhizophydiales Stephanodiscus 89.5\nOL869117 uncultured Zygophlyctidales Diatoma 89.5\nOL869120 uncultured Rhizophydiales Cosmarium 89.5\nOL869112 uncultured Zygophlyctidales Cyclotella 89.2\nOL869113 uncultured Lobulomycetales Fragilaria 87.9\nOL869114 uncultured Lobulomycetales Fragilaria 87.6\nOL869116 uncultured Rhizophydiales Synedra 87.0\nOL869121 uncultured Rhizophydiales Fragilaria 86.4\nOL869118 uncultured Polyphagales Yamagishiella 83.6\nFigure 1. Phylogenetic trees created on NCBI using (A) the 100 best BLAST matches and (B) sequences of chytrids known to infect phytoplankton (data obtained from Van den Wyngaert et al 2022). ASV 8 (shown in red) represents the most abundant fungal ASV in our sample (87% of Chytridiomycota-assigned reads).\nstarting on line 200: It is a bit unconventional to write a scientific results and discussion section with this Q1, Q2, et c formatting. I suggest making these individual paragraphs with section headers.\nResponse: Yes, we agree. Individual paragraphs with headers are now implemented.\nlines 212-234: The discovery of the fungal preference for the N2-fixing heterocyst and N-rich akinete storage cells over the vegetative cells is extremely interesting! A remarkable result indeed. This high nitrogen assimilation by the marine fungi associated with the N2-fixing heterocysts makes me wonder, if organic nitrogen assimilation is an important part of the ecology of marine fungi in general. It reminds me of similarly high organic N assimilation activity inferred for marine fungi in the Namibian upwelling zone (doi.org/10.1038/s41396-021-01169-5). Could the authors comment on organic N assimilation more generally as relates to marine fungal ecology in the discussion? And, also - what the implications of fungal organic N assimilations might be for the marine N cycle?\nResponse: Interesting thought. In filaments of Anabaena/Dolichospermum, N2 is converted to NH4+, and thereafter into glutamine and glutamate. Those amino acids are the principal export forms of fixed nitrogen from heterocytes to neighboring cells. Akinetes are differentiated resting cells, functioning as storage and survival structures. They accumulate organic nitrogen compounds, especially cyanophycin. The source of nitrogen in heterocytes and akinetes is thus mainly organic yet with different complexity—less complex amino acids in heterocytes and more complex cyanophycin in akinetes.\nIn general, marine fungi encode and express a diverse spectrum of carbohydrate- and protein-degrading enzymes in the oceanic water column (Baltar et al 2021, Breyer et al 2022). For instance, Orsi et al (2022) nicely demonstrated the direct assimilation of 13C-labelled org. carbon by diverse fungi in marine waters and sediments. Data on org. nitrogen assimilation, however, is scarce and also not included in the study by Orsi et al (2022). We thus believe that such discussion goes beyond the scope of our study—given that we focused on parasitic, early-diverging fungi (not covering the enormous diversity of marine fungi) and that we did not differentiate between different nitrogen forms.\nTo emphasize the role of fungal parasites on N cycling during cyanobacterial blooms, we added the conclusion: “We report on the significant yet largely overlooked role of fungal parasites in shaping the dynamics of N2-fixing cyanobacterial blooms. Our findings highlight that chytrid infections not only reduce the efficiency of N2 fixation but also alter the fate of newly fixed nitrogen, with up to 27% being channeled to fungal parasites.” (line 358 ff.)\nlines 309-311: This is a really interesting discovery.\nResponse: Yes, we were also surprised that fungal infections on Baltic Sea cyanobacteria have gone mostly unnoticed so far, given that the Baltic Sea is one of the best monitored and studied water bodies globally. We believe that this discovery will change our understanding of control mechanisms, trophic transfer, and biogeochemical cycles during algal blooms.\nlines 328-329: This is where the phylogenetic tree of the 18S gene from the chytrids studied her would be useful, R crass and R anabaena should also be represented in the tree. Is it the same species or different? lines 331-332: This can be easily resolved using a simple 18S phylogenetic tree analysis.\nResponse: We thank the reviewer for this comment. In our dataset, 97% of fungal reads belonged to the class Chytridiomycetes (Fig. 2A below), and ASV_8 alone represented the vast majority of those reads (Fig. 2B). Moreover, most of the remaining chytrid-related ASVs shared at least 99% identity with ASV_8 (Table 2). At this level of sequence similarity, it seems not possible to distinguish different species with confidence.\nThis limitation arises because the sequenced marker—the V7/8 region of the 18S rRNA gene—offers typically genus- to family-level resolution in early-diverging fungi. Many distinct chytrid species share identical or nearly identical V7/8 sequences, and consequently, phylogenetic trees based on this region alone cannot separate species-level lineages.\nRobust discrimination among chytrid species generally requires additional loci (e.g., full-length 18S, 28S, and/or ITS) rather than short amplicons. We added this information to line 269 ff.\nSince our V7/8 sequences shared at most 92% identity with the 18S rRNA gene sequence of a zoosporic fungus infecting freshwater Dolichospermum akinetes, indicated as R. akinetum (accession no. OL869110 in Van den Wyngaert et al (2022)), we infer that the Baltic Sea taxa are distinctly different from those occurring in lakes (as now stated in line 273 ff.). Very recently, sequences of zoosporic fungi infecting phytoplankton have also been published from the Baltic Sea, published August 7, 2025 (Van den Wyngaert et al 2025), but the corresponding 18S sequences were not yet available in NCBI (last checked August 26, 2025).\nFigure 2. (A) 18S-based community composition. (B) Number of reads of Chytridiomycota-assigned ASVs, derived from our environmental Baltic Sea sample.\nTable 2. Percent identity of Chytridiomycota-assigned ASVs, using ASV #8 as a reference.\n# ASV_ID Percent Identity_to_ASV_8 Percent_reads\nReviewer #2 (Remarks to the Author):\nFeuring and colleagues present their study showing that fungal parasites significantly impact nitrogen cycling during cyanobacterial blooms by infecting key cell types in Dolichospermum, extracting substantial amounts of host-derived carbon and nitrogen. These infections divert newly fixed nitrogen to fungi, rivaling bacterial uptake, and suggest fungal parasitism is a widespread and influential factor in bloom dynamics in the Baltic Sea.\nThough prevalent in aquatic ecosystems, we still only have a limited understanding of chytrid ecology and studies like this are therefore critical. The team are experts in applying sophisticated isotope techniques to study chytrid-algal interaction and this approach is used well here.\nA generally well-written manuscript. I have a few points to consider.\nWe thank the reviewer for the encouraging evaluation of our manuscript and helpful suggestions, which improved our manuscript. Below, we provide a detailed, point-by-point response.\nLine numbers correspond to those in the final document without track changes. Additional edits were made to ensure compliance with the Nature Communications formatting guidelines (a title without punctuation, shortened abstract, etc.).\nTo what extent the novelty of the system needs stating more clearly. This a new marine observation but has it been shown to some extent already in freshwater ecosystems?\nWe clarify that fungal infections on pelagic cyanobacteria have been observed previously in freshwater (l. 61), and we detail those previous observations in the discussion (l 316) by comparing our observations with those made in freshwater. To our knowledge, we were the first to report fungal infections in Dolichospermum (formerly Anabaena) in a coastal system (Baltic Sea) when submitting our manuscript, but just very recently, on Aug 7th, 2025, Van den Wyngaert et al (2025) also report such infections on Dolichospermum in the Northern Baltic Sea (we included this reference in the revised version). The authors, however, neither focus specifically on Dolichospermum nor include any activity and transfer rates in their study. But their study confirms that fungal infections on phytoplankton are widespread across the Baltic Sea, underlining the high relevance of our novel, first-time quantitative data.\nTo what extent is this a local phenomenon for the Baltic Sea? Could a statement be made of the wider significance of this work?\nResponse: Fungal infections are not a local phenomenon. Infections have been reported worldwide, including freshwater and marine systems. We have included this wider significance statement in the introduction.\nLine 53 ff.: Mounting evidence, however, suggests that parasitic fungi, belonging mostly to the order Chytridiomycota (hereafter referred to as chytrids), are widespread in coastal (Gutiérrez et al 2016, Hassett and Gradinger 2016, Kilias et al 2020) and freshwater ecosystems (Sime-Ngando 2012, Van den Wyngaert et al 2022). While thriving on major phytoplankton groups, including diatoms, cyanobacteria, and dinoflagellates, fungal parasites parasitize up to 90% of the phytoplankton host population (Gerphagnon et al 2017, Kagami et al 2006, Rasconi et al 2012).”\nWhile we provide some exemplary references in the introduction, there are numerous additional references available from the literature, including e.g. those that report direct observations in coastal regions: (Garvetto et al 2019, Hanic et al 2009, Hassett et al 2017, Jobard et al 2010, Lepelletier et al 2014, Scholz et al 2014, Scholz et al 2016a, Scholz et al 2016b) (Cleary et al 2017), or those that observed a link between phyloplankton biomass and chytrid sequence reads (Duan et al 2018, Gao et al 2009) (Gutiérrez et al 2010, Wang et al 2018).\nThe early results on Dolichospermum distribution – is this not already known? This feels rudimentary.\nResponse: We assume that the “early results on Dolichospermum distribution” refer to the data shown in Fig. 1a.\nYes, the growth season of Dolichospermum lasting from mid-June to the beginning of August (with lower abundances occurring until the beginning of October) is known for the Southern Baltic Sea, given that this area is frequently monitored by the HELCOM and IOW-internal monitoring program (Kownacka et al 2020). The fact that we are here using publicly available monitoring data is stated in the legend of Fig. 1, line 647 ff., and in the method section, line 580/581. We included this data to align our own monitoring period, including fungal infections as a novel parameter, over the main growth season (as shown in Fig. 1b).\nThe selection of specific cell types is fascinating and a key result. Could you observe the rhizoids inside the cells? Do they stay in the targeted cell-type or move to the next?\nResponse: Good point. For fungal cells infecting heterocytes/akinetes, we did not observe rhizoids reaching out to the neighboring vegetative cells. We have now added this information in line 113 ff. By contrast, rhizoids infecting vegetative cells grew into the neighboring cells, often extending over several vegetative cells (also discussed in line 261 ff. and shown in Supplementary Fig. S4i, j).\nEncysted zoospore feels contradictory. Can you define? E.g. flagellum retracted?\nResponse: For clarity, we revised our terminology as “zoospore encystment”, and specified this stage as follows: “zoospore loses the flagellum, secretes a wall, and adheres to the host cells” (lines 151 f.). A visual example is shown in Fig. 3a.\nThe observation of more bacteria associated with infected cells. Brilliant result. Could more be explained on this?\nResponse: Yes, we agree. Filamentous N2-fixing cyanobacteria are commonly efficient in retaining newly fixed nitrogen, and they are embedded in a mucilage sheath that acts as a physical and chemical barrier, both limiting bacterial colonization. We hypothesize that those protective properties are deteriorated due to fungal attacks and related fungal enzymatic activities. We have included this information in the discussion (line 341 ff.).\nI am not familiar with isotopic techniques used, so cannot comment on the validity of the approach.\nResponse: It may not be necessary, but we would like to add some words on the validity of our approach.\nIsotope tracing at the single-cell level using secondary ion mass spectrometry (SIMS) is widely recognized as one of the most advanced approaches in environmental microbiology for quantifying microbial activity and tracing elemental transfer within tightly linked microbial assemblages (e.g., Mayali 2020, Pett-Ridge and Weber 2022). Our laboratory has long-standing expertise in this technique, developed through years of research and multiple studies (e.g., Klawonn et al 2016, Klawonn et al 2019, Klawonn et al 2021). We are therefore confident in the validity and robustness of our isotopic tracing approach."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The Reviewers have done a nice job addressing my comments, the only last remaining issue is that the nice figures and analyses that the provided in the rebuttal document were not added to the main text figures or supplemental for the revised manuscript. These new phylogenetic analyses (in particular the tree) should be reported in the manuscript (they could be in the supplemental). If the authors can make this change, then I would recommend publication."}, "author_response": "We agree and have added the phylogenetic tree to the Supplementary (Supplementary Figure S5).\nWe wish to thank the reviewer again for the careful review."}]

Decision letter