Significance Atmospherically transported dust from the Saharan desert provides pulses of biologically important nutrients, including iron, to ocean surface waters. The biological response to these ephemeral events is not fully known, especially among the heterotrophic microbial community. Here we use the well-characterized Vibrio genus as a model for heterotrophic bacterial response. We demonstrate that Saharan dust nutrients, deposited in tropical marine waters, can promote Vibrio bloom formation and suggest that dust-associated iron is an important driver of Vibrio population dynamics. This work shows not only the role of fast-acting heterotrophs in the biogeochemical cycles of environmental pulses of iron, but it also highlights an important factor in the growth of bacteria that can cause disease in humans and marine organisms.

Abstract Vibrio is a ubiquitous genus of marine bacteria, typically comprising a small fraction of the total microbial community in surface waters, but capable of becoming a dominant taxon in response to poorly characterized factors. Iron (Fe), often restricted by limited bioavailability and low external supply, is an essential micronutrient that can limit Vibrio growth. Vibrio species have robust metabolic capabilities and an array of Fe-acquisition mechanisms, and are able to respond rapidly to nutrient influx, yet Vibrio response to environmental pulses of Fe remains uncharacterized. Here we examined the population growth of Vibrio after natural and simulated pulses of atmospherically transported Saharan dust, an important and episodic source of Fe to tropical marine waters. As a model for opportunistic bacterial heterotrophs, we demonstrated that Vibrio proliferate in response to a broad range of dust-Fe additions at rapid timescales. Within 24 h of exposure, strains of Vibrio cholerae and Vibrio alginolyticus were able to directly use Saharan dust–Fe to support rapid growth. These findings were also confirmed with in situ field studies; arrival of Saharan dust in the Caribbean and subtropical Atlantic coincided with high levels of dissolved Fe, followed by up to a 30-fold increase of culturable Vibrio over background levels within 24 h. The relative abundance of Vibrio increased from ∼1 to ∼20% of the total microbial community. This study, to our knowledge, is the first to describe Vibrio response to Saharan dust nutrients, having implications at the intersection of marine ecology, Fe biogeochemistry, and both human and environmental health.

Bacteria in the Vibrio genus are globally distributed in marine environments but typically make up a minor portion of the total microbial assemblage (1, 2); however, Vibrio have been shown to bloom in response to often poorly characterized environmental factors (3, 4). Like other opportunistic heterotrophic bacteria, Vibrio can have disproportionately large impacts on carbon and nutrient processing because of their ability to reproduce rapidly and respond to pulses of newly available resources (2, 5⇓–7). Characterized as “opportunitrophs,” Vibrio have a broad genomic and metabolic repertoire (8), allowing them to compete in highly variable nutrient environments ranging from the open ocean to pathogenic associations with animal hosts (3, 9). This genus includes many well-known pathogens of marine organisms and humans, and disease incidence has risen sharply in the last 20 y (10, 11). Common human pathogens include the causative agent of the severe diarrheal disease cholera (Vibrio cholerae), shellfish-associated gastroenteritis (Vibrio parahaemolyticus and Vibrio vulnificus), and seawater-associated wound infections (Vibrio vulnificus and Vibrio alginolyticus) (10). Studies examining the environmental drivers and distribution of Vibrio have largely focused on the role of Vibrio in disease, generally overlooking the importance of Vibrio in the biogeochemical cycling of key nutrients and trace metals (3, 4). Iron (Fe) is an essential micronutrient for Vibrio growth in the environment as well as during host invasion, where it is actively sequestered by the host to prevent bacterial colonization (9). Vibrio have evolved to be adept scavengers of Fe in a variety of conditions (12). Despite the importance of Fe for growth, there has been little characterization of the effects of environmental Fe enrichment on Vibrio population dynamics and the role of these bacteria in Fe cycling in marine systems.

Fe can be a limiting micronutrient in marine primary and secondary production (13, 14). As an essential cofactor in many metabolic processes—including aerobic respiration, photosynthesis, and nitrogen fixation—its availability can be a determinant in the cycling of carbon (C) and biologically important macronutrients, like nitrogen (N) and phosphorus (P) (13, 15, 16). Dissolved Fe (dFe) is believed to be the most biologically available fraction of Fe, but is present in vanishingly low amounts in marine systems around the world, especially due to the low solubility of Fe(III) in seawater (17). Heterotrophic bacteria, including Vibrio, play a key role in Fe cycling (14, 15, 18, 19), in part by modulating Fe solubility through secretion of high-affinity Fe-chelating siderophores (20, 21). Fe-siderophore complexes allow active uptake into the bacterial cell (20) and provide a usable exogenous source of Fe for phytoplankton (22, 23). Although most studies of Fe enrichment in marine systems have focused on autotrophs (24, 25), heterotrophic bacteria have been shown to have a higher Fe per biomass quota than many phytoplankton (18), accounting for up to 80% of the total planktonic uptake in some systems (26).

Atmospheric dust deposition is a major source of Fe in the global ocean (27) and is estimated to deliver more than triple the amount of dFe as riverine inflow (28). Globally, the Saharan desert is the largest source region of this atmospheric dust–Fe (27). Dust originating in northern Africa is swept across the Atlantic Ocean along easterly trade winds, producing spatiotemporal gradients of dust deposition in the Caribbean and southeastern United States, especially during the summer months (29, 30). Increased atmospheric processing time, associated with long-range transport as well as wet deposition (rain washout), are hypothesized to alter components of atmospheric dust and produce a soluble and highly biologically available form of Fe (17, 27). Although the response of marine autotrophs to dust deposition has been investigated (15, 25), the full biological response to the episodic deposition of dust–Fe to ocean surface water, especially among microbial communities, has yet to be clearly elucidated. Evidence is growing that heterotrophic bacteria, especially among the class γ-proteobacteria, may play a role in processing deposited minerals and nutrients (31, 32), but studies to date, which have largely focused on bulk bacterial response and longer timescales, have shown equivocal results (33, 34). We hypothesized that specific opportunistic bacteria, like Vibrio, can respond quickly to newly available dust-associated Fe and suggest a role of dust–Fe as a driver of Vibrio population dynamics.

Results Vibrio Response to Simulated Saharan Dust Deposition in Marine Surface Water. To determine the effect of Saharan dust nutrients on Vibrio growth, source material from the Saharan desert (Morocco) was added to microcosms containing natural unfiltered seawater collected in the Florida Keys (US). Source material (characterization shown in Tables S1 and S2) was manipulated to simulate effects from long-range atmospheric transport and wet deposition and is referred to as Dust SIM (SI Methods). Microcosms included surface water collected over multiple dates from three sites for each experiment, representing a cross-shelf gradient (onshore, near shore, and offshore (at Looe Key Reef in the lower Florida Keys)] (Fig. S1). A broad range of dust deposition scenarios (20 μg⋅L−1 to 30 mg⋅L−1) were simulated. The lowest Dust SIM addition (20 μg⋅L−1) increased dFe in seawater to 1.19 nM (±0.08 SEM) above the background levels of 0.90 nM (±0.04 SEM) found in nonamended surface water (Table S3). This addition provided a small but significant increase (P < 0.05), consistent with in situ observations of dFe during dust events (Table S3; also see Fig. 4). Table S1. Analysis of Moroccan source material Table S2. Mineralogical makeup and size analysis of Moroccan source material (% of total) Fig. S1. Sampling locations for Dust SIM seeded experiments (Florida Keys): inshore (500 m from Mote Tropical Research Lab, Summerland Key, FL), near shore (1 km from shore), and offshore (10 km from shore), Looe Key Coral Reef. Table S3. Analysis of dFe in Florida Keys seawater (July 2014) After the Dust SIM amendments to seawater, the growth of total culturable Vibrio on a selective medium was normalized across experiments by calculating the ratio of colony-forming units (cfu) per milliliter at 24 h to those at 0 h. All Dust SIM additions, except the 40 μg⋅L−1 addition (due to high variability among replicates), had significantly higher growth compared with nondust controls, and the response was largely dose-dependent (Fig. 1; Table S4). At the lowest Dust SIM addition (20 μg⋅L−1), mean growth was six times as great as that for nondust controls (49.0 ± 19.6 SEM and 8.5 ± 3.6 SEM, respectively, P < 0.001). Neither the location (inshore, near shore, or offshore) nor the date of sampling had a statistically significant effect (P > 0.05). Fig. 1. Florida Keys surface water Vibrio response to a range of Saharan Dust SIM additions. Vibrio growth was normalized among experiments by comparing the ratio of cfu/mL 24 h after Dust SIM addition (T 24 ) to time 0 (T 0 ) for [n] replicates (mean ± SEM). Asterisks indicate those amendments in which growth was significantly different from the no-dust (0 mg⋅L−1) control (mixed linear model; Tukey’s post hoc test). *P < 0.05; ****P < 0.0001. Additional data are shown in Table S4. Table S4. Total Vibrio concentrations from natural seawater community amended with increasing concentrations of Dust SIM Vibrio Utilization of Dust-Associated Fe. To evaluate the specific effects of Fe in Saharan dust on Vibrio growth, cultures of individual strains were grown in a novel, Fe-limited seawater (Vib-FeL), which was replete in key macronutrients (N and P) and carbon substrate and supplemented with all essential trace elements needed for growth, except for Fe (SI Methods). The relative growth of individual strains of V. alginolyticus and V. cholerae (identified as dust-responsive in initial experiments) were tested in three separate culture conditions: (i) Vib-FeL alone, (ii) Vib-FeL with Dust SIM (providing 0.89 μM Fe), and (iii) Vib-FeL with FeCl 3 (providing 4.37 μM Fe), as a positive control. After overnight incubation (18–24 h), the proportional growth was compared for all treatments (Fig. 2; Table S5). Vib-FeL alone restricted growth of both test strains. The addition of Dust SIM led to significant growth of both V. cholerae and V. alginolyticus compared with nondust controls (13 and 18 times as great, respectively; P < 0.05). The proportional growth for the Dust SIM amendments was not significantly different from that noted for the FeCl 3 controls, which had 20 and 23 times as much growth for V. cholerae and V. alginolyticus, respectively, as did nondust controls (Fig. 2; Table S5) and confirmed that Dust SIM could alleviate Fe limitation in this medium. Fig. 2. Vibrio spp. response to Fe and dust. Growth of V. alginolyticus (n = 3; A) and V. cholerae (n = 2; B) in Fe-limited seawater (Vib-FeL) alone, amended with 4 μM FeCl 3 (Fe), and Dust SIM (Dust) containing 0.89 μM Fe. Vibrio growth was normalized among experiments by comparing the ratio of cfu⋅mL−1 after Dust SIM addition (T n ) to time 0 (T 0 ) (mean ± SEM). Asterisks indicate significantly greater growth with the addition of Fe and dust compared with growth in Vib-FeL alone (ANOVA). *P < 0.05; **P < 0.01; ***P < 0.001. There was no significant difference between treatment with Dust SIM and treatment with Fe. Additional data are shown in Table S5. Table S5. V. alginolyticus and V. cholerae concentrations in minimal medium (Vib-FeL) amended with FeCl 3 and Dust SIM Fe concentrations across dust deposition events are highly variable (27, 35); therefore, we evaluated growth of V. alginolyticus in Vib-FeL across a range of scenarios, providing additional dFe at concentrations from 5 to 836 nM. All Dust SIM additions increased the proportional growth of V. alginolyticus (Fig. 3; Table S6). The lower additions of 5, 10, and 21 nM dFe resulted in approximately four times the growth of no-addition controls. The higher dFe amendments, 201 and 836 nM, resulted in growth that was 8 to 11 times that of controls (P < 0.001), respectively (Fig. 3; Table S6). Fig. 3. Response of Vibrio alginolyticus to a range of Dust SIM dFe concentrations in Fe-limited artificial seawater (Vib-FeL). Vibrio growth was normalized among experiments by comparing the ratio of cfu⋅mL−1 of V. alginolyticus 24 h after Dust SIM addition (T 24 ) to time 0 (T 0 ) (n = 3; mean ± SEM). Asterisks indicate those amendments in which growth was significantly different from the no dust addition (0 Dust SIM ) control (ANOVA) *P < 0.05; ***P < 0.001. Additional data are shown in Table S6. Table S6. Vibrio alginolyticus concentrations in minimal medium (Vib-FeL) amended with increasing concentrations of Dust SIM Vibrio Growth in Response to Natural Saharan Dust Events. Offshore sampling sites at Looe Key Reef, in the lower Florida Keys (US), and Ragged Point, Barbados, were chosen based on a 30-y dataset of atmospheric dust sampling, demonstrating strong seasonal pulses of African dust arriving at these locations almost entirely in the summer months (June through September) (29, 30). Satellite and aerosol-modeling products were used to monitor evolution of individual dust events from the coast of Africa (typically every 5–10 d, July through August) and their transit across the Atlantic (Fig. 4). Field collections were conducted in the Florida Keys (summer 2013 and 2014) and Barbados (summer 2014) during three separate Saharan dust events (SDEs). Fig. 4. Vibrio response to natural SDEs. (Left) Vibrio concentrations (cfu⋅mL−1) (n = 3; mean ± SEM) in surface waters during the arrival (shaded bar) of SDE and 3–5 d after dust arrival in Barbados (A) and the Florida Keys (B and C). Dashed lines represent baseline surface water Vibrio concentrations (348 ±122 SEM, n = 3 for Barbados and 45.4 ± 2.5 SEM, n = 42 for Florida), determined during the dust season (April– August) but not associated with a SDE. For each SDE, Vibrio concentration on the date of peak response was significantly different from all other dates. *P < 0.05; ***P < 0.001. Double hash indicates nonconsecutive sampling date (C). Mean (± SEM, n = 3) dFe concentrations were determined for A and B only; bold values indicate significantly lower dFe than that observed on the dust arrival date (ANOVA P < 0.05). (Right) Modeled dust aerosol depths (Naval Research Laboratory (www.nrlmry.navy.mil/aerosol/) (study site indicated by red circle). In July 2014, 1 wk before the arrival of dust, dFe in surface waters in the lower Florida Keys was 0.90 nM (±0.08 SEM), increasing to 3.24 nM (±0.04 SEM) with the arrival of a SDE (P < 0.001) (Table S3). Similarly, the highest dFe during the August 2014 study period in Barbados (2.22 nM ± 0.20 SEM) was measured during the arrival of a SDE. dFe levels were not measured in 2013. Within 14–24 h of the arrival of SDE (and concomitant dFe increase), surface water concentrations of culturable Vibrio increased significantly over background levels in all three sampling campaigns at levels ranging from 5 to 30 times the nondust levels (Fig. 4). At the peak of the Vibrio blooms in 2014, a 1.6- and 3.6-fold decrease of dFe was measured in Barbados and Florida, respectively (Fig. 4). Vibrio bloom conditions were transient, and levels returned to baseline within 24–48 h of the measured peak. As the Vibrio bloom expired, an increase of dFe was measured, followed by a tapering decline in dFe over the following 1–4 d to ∼1.6-fold decline in dFe compared to dust arrival values. Bacterial Community Composition Changes in Response to SDE. As part of a co-occurring study at Looe Key Reef, microbial community data were obtained for an additional four surface water samples between August 2011 and July 2013 using Illumina-based sequencing of the community 16S rDNA. Three of the samples were collected outside of a dust event. The final sample coincided with the dust event captured in July 2013. The abundance of operational taxonomic units in the Vibrio genus was significantly greater during this event than during all other sampling dates (P < 0.001). The relative abundance of Vibrio increased from <1.4% during nondust conditions to 19.8% of the total bacterial community during the 2013 SDE (Figs. S2 and S3). This increase was also confirmed by Vibrio culture counts and cell equivalent counts [quantitative PCR (qPCR)] (Table S7). The relative abundance of other genera including Pseudoalteromonas and Acinetobacter, belonging to the same class of bacteria as Vibrio (γ-proteobacteria), also increased significantly, whereas Pelagibacter declined (P < 0.05) (Fig. S3). Fig. S2. Bacterial community response to SDE in situ. (A) Composition of the bacterial community in surface water by date at Looe Key reef, FL, by using 16S rDNA bacterial community analysis. Vibrio are denoted by * (too low for the first three dates to be discernable on the figure). (B) Genera represented in bacterial community analyses. Fig. S3. Results of ANOVA for responsive bacterial taxa among dust and nondust periods in the Florida Keys. Proportions of each genus were square root arcsine transformed to approximate a normal distribution and compared by sample date (Aug 2011, Feb 2012, and Feb 2013 were nondust dates; July 2013 coincided with a SDE). For Vibrio, Acinetobacter, and Pseudoalteromonas, relative abundance in July 2013 exceeded all other sample dates, whereas the relative abundance of Pelagibacter was reduced during the July 2013 dust event. Table S7. Comparison of Vibrio abundance quantified by culturing on selective TCBS agar and Vibrio-specific qPCR

Discussion There has been considerable interest in the biological response of marine surface water communities to dust deposition, but results to date on the role of heterotrophic microorganisms have been equivocal. These recent investigations of heterotrophic response to dust-associated nutrients as well as the direct effect of Fe addition to marine waters generally measured bulk parameters (such as total bacterial abundance, bacterial respiration, or bacterial production), which could potentially fail to recognize community shifts and mask the emergence of functionally important taxa that can rapidly respond to nutrients (19, 31, 36). Additionally, studies that did assess community structure in response to dust typically focused on ≥48 h after dust addition, which could miss taxa that mount a more rapid response (31, 36). Finally, much of the recent work in this area has examined response in experimental microcosms and mesocosms (15, 31, 36), with few studies able to capture in situ response to natural dust events. In this study, evidence from both experimental manipulations and in situ time course observations demonstrated that opportunistic heterotrophic genera like Vibrio mount a robust and rapid growth response to dust addition and potentially play an important, but currently largely unexplored, role in the cycling of Fe. Fe acquisition is essential for Vibrio fitness and survival, both in the environment and in a host during infection (9, 12). V. cholerae, which is arguably the most studied of the Vibrio species and has a very well characterized genome, has >50 Fe acquisition genes spread across its two chromosomes for ferrous, ferric, and biologically complexed Fe uptake (12). This genetic repertoire suggests that Vibrio are effective competitors for Fe in many different environmental niches. The availability of Fe can also control the metabolism of other nutrients such as N and P, and can limit C utilization, effectively setting limits on growth for both open ocean and coastal heterotrophic bacteria (16, 33, 34). Potential colimitation is especially important in light of the fundamental contribution of heterotrophic bacteria to the cycling of C in marine ecosystems (33, 37). Vibrio and other rapidly responding heterotrophs likely play a pivotal, but as yet largely unexplored, role, in coupling Fe flux with C cycling in marine food webs, ultimately influencing global climate processes (18, 19, 37). In this study, we demonstrated that Vibrio rapidly exploit Fe and potentially other nutrients provided by Saharan dust, driving a significant, but temporary, population bloom. These blooms, which were observed within 24 h of dust arrival, also likely preceded stimulation of autotrophic picoplankton and other phytoplankton, which have previously been shown to take ≥48 h to respond to dust or Fe inputs (e.g., ref. 31). This finding suggests that Vibrio blooms were not due to increased organic matter production from autotrophs, but were responding directly to the dust inputs, which is also supported by the microcosm studies using monocultures of Vibrio. Furthermore, the data suggest that dust–Fe can support blooms across a wide range of dust and Fe deposition scenarios. Vibrio growth in natural seawater continued even at the highest Dust SIM –Fe additions (836 nM), suggesting increasing Fe limitation, which was unexpected (34). Although the bioavailable fraction of Fe added to the microcosms was likely reduced because of rapid oxidation and precipitation due physiochemical factors in natural seawater (17), Vibrio physiology suggests a high Fe demand with up to 5 μM needed for replete growth (12). Growth rates in Vibrio are among the fastest known of any bacteria, with doublings every 8–9 min recorded in some species (5). Additionally, several Vibrio species are able to fix N, a process that has a very high Fe demand (38, 39). Dust additions also likely added other limiting nutrients in addition to Fe, especially phosphate, which may help to explain the continued response even at very high levels (16). Finally, although the focus of this work was on Vibrio as a model for dust response, the relative abundance of other taxa (e.g., Pseudoalteromonas and Acinetobacter) also increased significantly with dust events and likely competed with Vibrio for uptake of additional Fe and nutrients. During in situ time course observations, Vibrio showed a robust bloom within 24 h of the arrival of SDE and concomitant high dFe concentrations. At the peak of the bloom, Vibrio species increased their proportion in the community by more than an order of magnitude, relative to nondust conditions. These results indicate that this is not an exogenous import of bacteria, but rather an autochthonous response to the addition of biologically necessary nutrients like dFe. Although it has been reported that some bacteria can travel associated with dust aerosols, genus-level profiling of Saharan dust has not revealed the presence of Vibrio (40). During the 2014 field studies, the drawdown of dFe to the predust level of 0.9 nM (Fig. 4 and Table S3) at the peak of the bloom also indicated utilization of the majority of introduced dFe by blooming Vibrio and other responsive γ-proteobacteria (31). As a genus, Vibrio is one of the most highly investigated groups of environmental microbes. Aided in part by its ease of culturability (3), Vibrio is an excellent candidate model to examine opportunistic responses and roles in larger ecosystem functioning (8). Vibrio, which typically comprise <1% of the ocean surface bacterioplankton community (2), could be considered conditionally rare taxa (CRT) (41). CRT are subject to dramatic blooms, potentially playing an outsized role in the ecology of a system (1, 41, 42). Vibrio have shown explosive growth in response to nutrient enrichment (6, 7). This adaptive feast-or-famine life strategy allows exploitation of spatially and temporally variable resources, leading to bloom conditions. This strategy also subjects Vibrio to kill-the-winner top-down controls, such as grazing and viral lysis, both of which tightly control γ-proteobacteria and Vibrio populations (7, 43, 44). The population declines observed in the field 24–48 h after SDE-induced Vibrio blooms could be attributed to these top-down control pressures. The bloom decline corresponded with a spike in dFe concentrations, supporting the notion of lysis and bacterivory in the release of dFe, warranting further investigation. The ecosystem consequences of such a large turnover of dFe on biogeochemical cycles also remains to be determined. Bacterial grazing experiments have demonstrated that 90% of Fe remains in the dissolved fraction after 24 h (14). Additionally, Fe released by viral lysis of heterotrophic bacteria (as demonstrated in V. alginolyticus) is highly bioavailable to the plankton community, including autotrophic diatoms, with the bioavailability of this released Fe exceeding that of siderophore-bound Fe (21). This dFe is capable of supporting up to 90% of primary production in some systems (45). Taken together, the bloom-bust cycle of Vibrio population growth in response to Saharan dust could have an important role in trophic transfer of labile Fe to primary producers. Vibrio can overcome predation pressure by forming close associations with marine organisms and plankton (46). In this work, the population bloom of Vibrio appeared transient, but further investigation is needed to determine whether planktonic-associated hot spots exist after bloom termination. The potential for particle attachment is particularly salient in the case of disease-causing species like V. cholerae or V. alginolyticus, which have both been shown to be highly responsive to dust–Fe. These species are known to associate with zooplankton and marine organisms, directly influencing their survival in the environment and routes of transmission to potential hosts (3, 9, 46). This study, to our knowledge, is the first to demonstrate that Vibrio species, as a model of a rapidly responding opportunistic heterotroph, are highly responsive to Saharan dust–Fe and associated nutrients and indicate a role for these bacteria in processing dust–Fe in marine ecosystems. In particular, these results suggest that otherwise rare heterotrophic bacterial taxa are among the first responders to introduced Fe and other nutrients from dust and likely precede responses by autotrophs. The Sahara and Sahel are particularly vulnerable to further drying due to changes in climate and land-use patterns (47), potentially increasing dust export from this region. Coupled with the fact that dust–Fe fertilization of marine systems has been suggested as a driver of past paleoclimatic change (48), a mechanistic understanding in the modern ocean is critical to making predictions about future oceanic production and climate scenarios. The discovery of dust–Fe as a factor in Vibrio population dynamics is an important first step that warrants further investigation to inform future predictions about Vibrio-related disease and Vibrio impacts on global biogeochemical cycles.

Methods Dust SIM Experimental Additions. The US Geological Survey (USGS) provided mineralogical and elementally characterized Saharan source material (SI Methods) collected from a highly weathered dune in Morocco (exact location was not specified). Saharan source material was manipulated in the laboratory to simulate atmospheric processing and wet deposition and is referred to as Dust SIM (SI Methods). Natural surface water from the Florida Keys and artificial Vib-FeL microcosm experiments were seeded with a broad range of Dust SIM additions because of the high spatial and temporal heterogeneity of dust deposition in any single event, as well as the high variability in dust concentrations resulting from rain washout, which is the dominant mode of deposition in Florida (∼80%) (29). To quantify Dust SIM loadings for experimental additions, Dust SIM was added [1:150 (vol/vol)] to 5 mL of ultrapure Milli-Q water by using trace metal clean chemistry techniques (49), followed by further 1:2 serial dilutions in Milli-Q water. dFE (<0.2 μm) was measured by using inductively coupled plasma MS (ICP-MS) elemental analysis (SI Methods) for triplicate samples of each dilution. The Fe content of trans-Atlantic transported dust has been measured to be close to the upper crustal value of 3.5% (29, 30); we therefore quantified Dust SIM amendments by Fe content to allow for an approximate calculation of dust loading (mg⋅L−1) for each addition, considering a fractional solubility of the Moroccan source material to be 4.39% (SI Methods). Dust SIM additions for seeding experiments were calculated to be: 0.02, 0.04, 0.18, 0.37, 0.78, 7.3, and 30 mg⋅L−1 of dust (Table S8). The higher values include representative amounts of deposition in regions closer to African source areas (e.g., Mediterranean), and the lowest values were environmentally relevant for the Caribbean and Florida Keys (e.g., during a wet deposition event) (29, 35). Table S8. Analysis of dFe in Dust SIM and calculated equivalent dust loading In microcosm experiments using natural seawater communities, surface water from the upper 1 m of the water column was collected in sterile trace-metal clean polypropylene bottles at sites in the lower Florida Keys (Fig. S1). Dust SIM solutions for seeding experiments were prepared as described above and added to replicate natural seawater microcosms. Microcosms were mixed and held at 30 °C for 24 h. Vibrio counts at the time of seeding (0 h) and after 24 h were determined by culture (cfu) on selective thiosulfate-citrate-bile salt-sucrose agar (TCBS; Oxoid), in triplicate. Additional details are provided in SI Methods. Fe-Limited Seawater (Vib-FeL) Experiments. To investigate the specific effect of Fe from Saharan dust on Vibrio growth dynamics, experimentation was done in a Fe-limited medium, referred to as Vib-FeL (SI Methods). Frozen cell stocks of V. alginolyticus [American Type Culture Collection (ATCC) strain 33839] and V. cholerae (ATCC 39315) were recovered in sterilized artificial seawater amended with 1% peptone and 0.5% yeast extract (ASW+PYE). Recovered cells, incubated for 12 h at 30 °C, were subcultured in 5 mL of fresh ASW+PYE (1:100 dilution) and allowed to grow to log phase as monitored by optical density at 600 nm (OD 600 ) on a spectrophotometer. Cells were washed twice and resuspended in Vib-FeL to eliminate Fe carryover. The experiment was initiated at time 0 by inoculating washed cells (1:100) into 5 mL of the appropriate culture conditions: Vib-FeL alone; Vib-FeL with FeCl 3 ; Dust SIM added to Vib-FeL (1:150 vol/vol); and incubated for 24 h at 30 °C on a shaking platform to maintain aeration. Vibrio abundance was determined by spread plating aliquots from each culture onto ASW+PYE agar plates at times 0, 18, and 24 h. Spread plates were incubated at 30 °C for 24 h, and cfu were enumerated and compared. Because dust deposition is highly variable (35), we also evaluated growth of V. alginolyticus (following the same cell preparation methodology) across a broad range of Fe-deposition scenarios by diluting Dust SIM as described above into Vib-FeL, providing dFe at concentrations from 5 to 836 nM dFe (Table S8). In Situ Response to Natural SDEs. SDEs were monitored by using satellite and modeling products from NASA (https://worldview.earthdata.nasa.gov/) and the Naval Research Laboratory (www.nrlmry.navy.mil/aerosol/), allowing for virtual real-time monitoring of SDE arrival and passage at two study sites: Ragged Point, Barbados (easternmost point of the island) (13.1667° N, 59.4333° W) and Looe Key Reef, FL (24.5475° N, 81.4067° W). Samples were collected within 24 h of the arrival of dust (Fig. 4). All surface water samples were collected in triplicate and kept at ambient temperature until processing (within 1 h). Sampling collection continued at 24-h intervals for 3–5 d after the modeled arrival of dust. Vibrio abundance was determined by spread plating each replicate surface water sample on Vibrio-selective TCBS medium, in triplicate. Spread plates were incubated at ambient temperature (∼30 °C) for 24 h, and cfu were enumerated and compared. Background nondust associated summer (May-August) Vibrio levels at Looe Key were determined by routine sampling between 2012 and 2014. Background levels at Barbados were determined in April 2013 (predust season). DNA was extracted from surface water samples collected from Looe Key Reef during nondust conditions and during the July 2013 SDE. Community DNA was sequenced by using Illumina MiSeq PE 250 sequencing of the V4 hypervariable region of the 16S rRNA gene (Tables S9–S11) (50) (SI Methods). Additionally, qPCR was conducted for total Vibrio counts from the extracted DNA (SI Methods). Field collections for SDE occurred in the summer rainy season, and isolated local showers were common; however, no precipitation data were available for the specific collection sites. Rain (very light) was only noted during or immediately before sampling at Barbados (11 and 12 August 2014). No rain was observed during sampling at Looe Key Reef. Table S9. Primers and Illumina adapters for the 16S rRNA gene hypervariable V4 region Table S10. Barcodes for reverse primers for 16S rDNA for sample tagging Table S11. Accession numbers for 16S rDNA microbial community sequence data All statistical analyses are described in SI Methods.

SI Results As part of a small subset of samples for in situ dust events and nondust events in the Florida Keys, Vibrio quantification methods (cfu vs. qPCR) were compared. Culture counts were primarily used in this study because of the specificity of TCBS medium, the ability to document changes in the community within 24 h in the field laboratory, and to allow isolates to be saved for additional work. However, given limitations in culture-based counts, a small experiment was conducted to compare results with cell equivalents as determined by Vibrio-specific qPCR (51). Contemporaneously collected samples from 6 separate days were analyzed (n = 4–6 per date); 2 d coincided with the 16S rDNA-based community analysis [Feb 2013 (nondust) and July 2013 (dust)], and the remaining 4 were collected during nondust periods (2013–2014). Samples were all collected at ∼2-m depth (midwater column) and were concentrated and extracted as described in Methods (for in situ community analysis). There were no significant differences between the Vibrio abundance estimated by culturing on selective TCBS agar and Vibrio-specific qPCR (Table S7). Additionally, there was no interaction between the fixed factors quantification method and sampling date (two-way ANOVA, P = 0.84), and differences between quantification method were not significant (P = 0.52). Vibrio abundance was significantly greater during the dust event (July 2013) compared with all other sampling dates.

SI Methods Saharan Source Material Analysis. The USGS provided Saharan source material collected from a highly weathered dune in Morocco (exact location was not specified). Elemental composition analysis was coordinated by the USGS and followed USGS Geochemical Landscape Project protocols (52). Compositional analysis of 37 elements was determined by ICP-atomic emission spectroscopy (ICP-AES) using USGS methodology (53) (Table S1). A Perkin-Elmer 2400 Carbon, Hydrogen, Nitrogen Analyzer (CHN) was used to determine the total carbon and N content in 91.21 mg of Moroccan source material (Table S1). Particle size was determined by using a Malvern Mastersizer 2000 laser analyzer, followed by textual classification according to USGS protocols (Table S2). Dust SIM Preparation. Atmospheric processing and wet deposition of Saharan dust were simulated in the laboratory by using Saharan source material. The material was manually dry-sieved through trace metal clean 335-µm mesh nylon followed by 150-µm mesh (Aquatic Eco-Systems Inc.) to remove coarse sand grains. To simulate high-altitude UV exposure and oxidize organic contaminants (including microbial contaminants), sieved material was exposed to UV irradiation (300 W Hg-lamp ∼254 nm) in a class 100 laminar flow hood in six 1-h increments with hand mixing of the material between exposures. Atmospheric acidification processes from anthropogenic aerosols (containing organic, nitric, and sulfuric acid species) are reported to render dust–Fe more soluble during atmospheric transport (17, 27). Dust passage and entrainment in the low pH environments of clouds and rain, as well as deposition by precipitation washout, were simulated by mixing 1 g of sieved UV-irradiated dust into 100 mL of 0.025 M HCl Optima (Fisher). HCl was used because seawater is already high in chloride, and results in minimal elemental alteration of the sample matrix during experimentation as well as provides maximal Fe dissolution. To eliminate large particulates that might promote adhesion and particle-supported bacterial growth, coarse material was allowed to settle out of the suspension, and the top 90 mL of the mixture was removed and filtered through an acid-washed, Milli-Q-rinsed, 0.2-µm pore-sized polycarbonate membrane (Whatman). The resulting stock material used in experimental manipulations was termed Dust SIM . The dFe fraction of Dust SIM that passed through the 0.2-µm membrane, considered the most bioavailable fraction (19), was 7.42 ppm (±0.12 SEM) as determined by ICP-MS. The fractional solubility of Moroccan source material Fe (16,900 ppm; Table S1) was calculated to be 4.39%. Dust SIM stock was confirmed to be free of culturable bacteria by direct spread plating of 0.1 mL of Dust SIM stock on experimental culture medium: ASW+PYE and the Vibrio selective media, TCBS (Oxoid). No growth was observed. ASW+PYE was prepared by dissolving Instant Ocean (Spectrum Brands) in Milli-Q water to a specific gravity of 1.023 and adding 1% Peptone (Fluka Analytical), 0.5% Yeast Extract (Oxoid), and 1.5% Bacto agar (Difco). ICP-MS Fe Analysis. All sample collection and trace-metal analyses were performed by using trace metal clean chemistry techniques (49). Samples were filtered through 0.2-μm track-etched polycarbonate filters and acidified to pH ∼1.8 (0.024 M HCl). Concentrations of dFe were determined by isotope dilution ICP-MS described in ref. 54. Natural SDE surface water samples were collected in acid-washed, 125-mL collection bottles just below the surface over Looe Key Reef, FL, and at Ragged Point, Barbados. Samples were frozen (within 30 min) for 1.5 mo before filtration to prevent Fe sorption to the bottle walls until sample analysis could be completed. Dust SIM Seeding Experiments in Surface Water Microcosms. Surface water, without filtration, was collected at sites in the lower Florida Keys (Fig. S1) and was used in seeded microcosm experiments. The offshore site was ∼10 km from shore, over the spur and groove reef at Looe Key Reef Sanctuary Preservation Area in the Florida Keys National Marine Sanctuary. The near shore (1 km from shore) and inshore (500 m from shore) sites were located near the Mote Tropical Research Laboratory on Summerland Key and were sampled by kayak. Grab samples of surface water were collected in the first meter at each sampling site in replicate experiments (March 2011, May 2012, and May 2013). Samples were collected in the late spring to avoid the influence of naturally occurring Saharan dust deposition, which is typically heaviest in the summer months (29). Samples were collected in sterile 1-L polyethylene bottles prepared by using trace-metal clean protocols (49). Each bottle was further rinsed three times with surface water at the collection site immediately before collection. Samples were transported to the laboratory in an ambient temperature cooler with a min-max thermometer, and experimental testing was initiated within 3 h of collection. No significant deviation from ambient temperature was observed in any experiment. Dust SIM dilutions were created in the same manner as described above in 5 mL of sample seawater from each site, followed by further 1:2 serial dilutions, providing calculated dust additions between 0.02 and 30 mg⋅L−1. A natural seawater (nondust) control was included for each site and experiment. Cultures were incubated for 24 h at 30 °C (a typical temperature for these sites during dust season) on a nutating mixer at 24 rpm (Fisher Scientific) to maintain aeration. Culturable Vibrio species were detected after spread-plating aliquots from each microcosm onto a Vibrio-selective medium, TCBS (Oxoid), in triplicate, at experimental times 0 and 24 h. Spread plates were incubated at 30 °C for 24 h, and cfu were enumerated per milliliter and compared. Fe Limited Seawater (Vib-FeL). To investigate the specific effect of Fe from Saharan dust on Vibrio growth dynamics, experimentation was performed in an Fe-limited medium, referred to as Vib-FeL, which was a modified version of the artificial saltwater medium, Aquil (55). Components of Vib-FeL were chosen with the lowest Fe content available. A solution of salts and nutrients was dissolved in Milli-Q water (18.2 MΩ × cm) containing NaCl (420 mM), MgSO 4 (2.5 mM), K 2 HPO 4 (0.1 mM), CaCl 2 (0.1 mM), NH 4 Cl (18 mM), and Na 2 PO 4 (42 mM). To remove potential contaminating Fe and other trace metals, the medium was passed through a column of Chelex 100 (Sigma) ion-exchange resin, according to the procedure of Price et al. (55), followed by microwave sterilization. After cooling (24 h), trace metals (50 µM EDTA-buffered solution of 2.5 × 10−8 M Co, 9.8 × 10−9 M Cu, 2.3 × 10−7 M Mn, 1.1 × 10−7 M Mo, and 3.9 × 10−8 M Zn) and a carbon source of sucrose (Sigma) [0.4% (wt/vol)] were added from 0.2-µm sterile-filtered stocks. All manipulations were performed in a laminar flow hood to minimize contamination. In Situ Community Analysis. Samples were obtained for community sequence analysis during three nondust periods [August 2011 (n = 7); February 2012 (n = 2); February 2013 (n = 3)] and during the 2013 SDE event at Looe Key Reef in the Florida Keys [July 2013 (n = 5)]. Water samples were collected with a sterile syringe (up to 60 mL) ∼1 m above the reef (∼2 m below the surface) and held on ice (<3 h). Aliquots (2 mL) were centrifuged at ∼13,000 × g for 20 min at 4 °C. The supernatant fluid was discarded, and the bacteria-containing pellet was stored at −20 °C. The protocol of Boström et al. (56) was used to extract environmental DNA from frozen pellets, with slight modifications. Lysis buffer [175 μL of 400 mM NaCl, 750mM sucrose, 20 mM EDTA, 50 mM Tris⋅HCl (pH 9.0)] and lysozyme (1 μL of 10 mg⋅mL−1) were added to the pelleted sample. After incubation at 37 °C for 30 min, proteinase K (100 μg⋅mL−1 final concentration) and SDS [1% (wt/vol) final concentration] were added, and tubes were incubated at 55 °C for 16–18 h. To aid in the precipitation of DNA, tRNA (50 μg) was used as a carrier molecule. Precipitation of DNA was initiated by adding 20 μL of 3 M NaAc and 120 μL of isopropyl alcohol and incubating for an hour at −20 °C. Samples were centrifuged (∼13,000 × g for 20 min), and the supernatant fluid was decanted, retaining the pelleted DNA in the original tube. Samples were then washed with 500 μL of EtOH (70%) and centrifuged (∼13,000 × g for 20 min), and supernatant fluid was discarded. A SpeedVac (Eppendorf Concentrator 5301) was used to dry the DNA pellet, which was then resuspended in 50 μL of TE (10 mM Tris⋅HCl and 0.1 mM EDTA, pH 8.0). Next-Generation Sequencing. The bacterial community was sequenced by targeting the hypervariable V4 region of the 16S rRNA gene using primers containing barcodes and Illumina adaptors (Tables S9 and S10) as described by Caporaso et al. (50). PCRs (triplicate for each sample) included 12.5 μL of Qiagen Taq PCR Master Mix, 0.5 μL of forward and reverse primers (2 μM final concentration), 2 μL of extracted DNA, and commercial sterile nuclease-free water added to a final volume of 25 μL. Amplified product (253 bp) was confirmed by gel electrophoresis; each band was excised and purified (MoBIO UltraClean GelSpin DNA Extraction Kit). PCR products were pooled and normalized into one sequencing sample based on the lowest sample concentration and mixed with 10% PhiX. Sequencing was completed with Illumina MiSeq PE250 (UGA Georgia Genomics Facility, Athens, GA). Sequence Analysis. Sequences were demultiplexed and barcodes were removed with the Illumina software (MiSeq Control Software Versions 2.2.0.2 and 2.3.0.8). Standard processing of the sequences was completed to remove sequences with poor quality and trim ends if under a quality score of 30, then only retaining sequences with at least 100 bp and 95% of bases with a quality score of ≥20 (57) (Gregory Hannon; hannonlab.cshl.edu/fastx_toolkit/index.htmL). The QIIME pipeline was used for all further sequence processing and data analysis, selecting default options (50). Mitochondrial sequences were removed by creating a corresponding sequence file from GenBank against BLAST sample sequences; matches to mitochondrial sequences with an e-value of ≥1−10 and 97% alignment were subsequently removed from analyses. Chimera Slayer was used to detect and remove chimeric sequences (58). Sequences that were not taxonomically classified within the bacterial kingdom were removed; these were Archaea, chloroplasts, and unclassified sequences. The remaining operational taxonomic units (otu) were picked by using an open reference frame, and taxonomy was assigned with the greengenes database (gg_13_5.fasta). The most abundant sequence for each otu, excluding singletons, was selected to create a representative set of sequences to align and create a phylogenetic tree (50). To analyze changes in the abundance of bacteria, proportions of each genus were square root arcsine transformed to approximate a normal distribution and compared by sample date using ANOVA (Fig. S3). Sequence data were uploaded to the NCBI BioProject database (accession no. PRJNA299413) (Table S11). Statistical Analysis. Estimates of growth were normalized among experiments by comparing the ratio of cfu⋅mL−1 at T n to that at T 0 , where T 0 represents time 0 and T n represents time n (e.g., 24 h) for individual microcosms. Results were presented as mean ± SEM. Proportional growth (T 24 /T 0 ) measurements from Florida Keys surface water amended with Dust SIM were first log transformed to meet normality assumptions. Statistical comparisons were performed with PROC MIXED by using a split-plot design with the factors of dust amendment, date, and location (SAS Version 9.3). Type III tests of fixed effects determined that neither location nor date, nor the interactions among effects were significant (P > 0.05). Dust amendments were compared by using post hoc pairwise comparisons. The Tukey–Kramer adjustment for multiple comparisons was used to control experiment-wise error rates, with P < 0.05 chosen to denote significant differences. All other statistical comparisons were made by using ANOVA and Tukey’s multiple comparison post hoc test (GraphPad Version 5.0.F). For all tests, significance was declared when P < 0.05.

Acknowledgments We thank Brian Hopkinson (University of Georgia) for consultation in developing Vib-FeL; Rebecca Auxier (University of Georgia) for ICP-MS analysis; Suzette Morman for analysis of Moroccan source material (USGS Crustal Geophysics and Geochemistry Science Center); and Kim Love Meyers (University of Georgia) for statistical consultation. We acknowledge collection support from the Mote Tropical Research Lab; Joseph Prospero (University of Miami); and Edmund Blades (Barbados Ministry of Health) for assistance in Barbados. This work was funded through National Science Foundation Grants EF-1015342 and OCE-1357423 (to E.K.L.) and OCE-1357140 (to W.M.L.); National Oceanic and Atmospheric Administration Oceans and Human Health Initiative S0867882 (to E.K.L.); the USGS Toxic Substances Hydrology Program (D.W.G.); and a student grant from the Association of Marine Laboratories of the Caribbean (to J.R.W.).

Footnotes Author contributions: J.R.W., D.W.G., and E.K.L. designed research; J.R.W., W.M.L., K.M.K., and E.K.L. performed research; J.R.W., A.M.E., W.M.L., J.L.J., and E.K.L. analyzed data; and J.R.W., W.M.L., D.W.G., and E.K.L. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Data deposition: The sequences reported in this paper have been deposited in the NCBI BioProject database, www.ncbi.nlm.nih.gov/bioproject (accession no. PRJNA299413).

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1518080113/-/DCSupplemental.