Table of Contents

• Coverage
• Dataset Description
  • Methods & Sampling
  • Data Processing Description
• Related Publications
• Parameters
• Instruments
• Project Information
• Funding

Strains and culture conditions:  All strains used in this study were taken from those used for a Long-Term Phytoplankton Evolution (LTPE) experiment (1). Prochlorococcus strains were streptomycin-resistant derivates of the high light-adapted strain MIT9312 obtained as described previously (2, 3), either before (Ancestor) or after 500 generations of evolution at either 400 ppm or 800 ppm pCO2 conditions (i.e., modern day or projected year 2100 conditions (4)). Alteromonas strains were derivatives of strain EZ55, originally isolated from a Prochlorococcus MIT9215 culture (3). As with our Prochlorococcus strains, we used both ancestral and evolved varieties of EZ55 co-evolved with Prochlorococcus at the two pCO2 treatments and subsequently isolated. Prochlorococcus cultures were revived from cultures cryopreserved with 7.5% DMSO in liquid nitrogen vapor, and Alteromonas cultures were revived from cultures preserved with 20% glycerol stored at -80o C. Prior to use in experiments, all Prochlorococcus cultures were grown in co-culture with Alteromonas EZ55 helpers (3) and were acclimated to culture conditions for at least 4 generations prior to data collection.

Alteromonas cultures were grown in YTSS medium (5) and Prochlorococcus cultures were grown in Pro99 medium (6) or PEv medium (1), both made in an artificial seawater base (ASW) (1). Prior to addition to co-cultures Alteromonas strains were pelleted at 2000 g for 2 minutes and washed twice in sterile ASW, then added to cultures at approximately 106 cells ml-1. Alteromonas was grown at 30o C with 120 rpm shaking. Unless otherwise noted, Prochlorococcus and co-cultures were grown in static 13 mL conical bottom acid-washed glass tubes under approximately 75 mmol photons m-2 s-1 cool white light in a Percival incubator set to 23o C. When medium additions were employed, all solutions were filter sterilized with a 0.2 mm filter. Cell densities of Prochlorococcus cultures to standardize inoculations between experiments were determined using a Guava HT1 flow cytometer (Luminex Corporation, Austin, TX) by the distinctive signature of these cells on plots of forward light scatter vs. red fluorescence (Fig. S1A). Day-to-day culture growth was tracked using the in vivo chlorophyll a module for the Trilogy fluorometer (Turner Designs, San Jose, CA) with a custom 3D-printed adapter designed for conical bottom tubes. Fluorometer measurements and cell counts were linearly related across the range of cells examined in this study (Pearson correlation coefficient 0.835, p = 1.38 x 10-6, Fig. S1B).

Concentration of Alteromonas exudates: EZ55 was grown in Pro99 media supplemented with 0.1% glucose to sustain growth in the absence of Prochlorococcus exudates. We scaled cultures up progressively from 12 mL to 2 L. The 2L culture was grown in a vented bottle with an outlet connected to a filter with 0.22 μm pore size. After removing most of the cells by centrifugation, we produced size-fractionated, concentrated exudates using tangential flow filtration using Sartorius Vivaflow 200 cassettes. The 2L culture supernatant was passed first through a 0.22 μm cassette using a Masterflex L/S peristaltic pump (Cole-Parmer) to remove bacterial cells, then through a 50 kDa module and a 5 kDa module in succession to produce >50 kDa and <50 kDa fractions that were each concentrated approximately 100-fold. A portion of the >50 kDa fraction was placed in boiling water for 5 minutes to denature proteins. When these concentrated extracellular products were added to culture media for growth experiments they were diluted 100-fold, returning them to approximately their original concentration prior to filtration.

Enzyme activities of exudates: We used a variety of enzymatic tests to evaluate the activity of Alteromonas exudates. We used the fluorescent conjugates 4-nitrophenyl-α-D-glucopyranoside, 4-nitrophenyl-β-D-glucopyranoside, and 4-nitrophenyl-N-acetyl-β-D-glucopyranoside to measure the activities of the glycolytic enzymes α-glucosidase, β-glucosidase and N-acetyl-glucosaminidase, respectively (7). Phosphatase activity was measured by following the hydrolysis of fluorescent substrate 4-methylumbelliferyl phosphate (8). Enzyme activity for these experiments was expressed as the rate of fluorescent product accumulation over the first 2-5 hours, minus the rate of accumulation in an exudate-free media control. Protease activity was measured after a 30 minute incubation based on Sigma’s non-specific protease activity assay using casein as a substrate (9). Siderophore activity was determined after a 30 minute incubation using chrome azurol S (10). Both fluorescence and optical density traces for these assays were measured using a Biotek Synergy H1 plate reader (Agilent, Santa Clara, CA). Hydroperoxidase activity was assessed by measuring the elimination of a hydrogen peroxide spike after 2 hours, tracking the hydrogen peroxide concentration using acridinium ester chemiluminescence via the injection protocol described previously (2) but modified for use with the Synergy H1. We also attempted to determine the role, if any, of intact EVs in enzyme function by sonicating a subsample of the >50 kDa fraction prior to assays. To accomplish this, the concentrates were treated with a Fisher Scientific sonic dismembrator for 3 cycles of 10 s with 10 s pause between (100W, 30% output efficiency). Both methylumbelliferyl and 4-nitrophenyl conjugate substrates are routinely used to measure membrane permeability due to their inability to cross membranes (11, 12), so an increase in apparent enzymatic activity post-sonication can be interpreted as an indication of membrane disruption.

ll statistical analyses were performed in R v. 4.4.1. Most analyses used linear models followed by post hoc extended marginal means testing of pairwise differences between treatment groups using the emmeans package (1). Assumptions of linear regression were checked for models by Shapiro-Wilk tests of the normality of residuals and plots of residuals vs. fitted values for homoscedasticity; where these assumptions were violated we used the Box-Cox procedure to find an optimal power transformation (2). Statistical differences between lysate and exudate protein localization counts were determined using Fisher’s exact test implemented in R.

NSF Award Abstract:
The function and stability of microbial communities in the ocean depends on exchanges of biological products and services between individual cells. Marine microbes are typically far apart from one another, so some of these exchanges occur through the release of products or services into the surrounding water, where they travel to other cells via simple diffusion. Understanding the degree to which such valuable products made by one organism are targeted to a specific partner, and how, has important implications for our understanding of the ecology and evolution of the marine microbiome. This project examines the role played by a poorly understood type of very small particle - extracellular membrane vesicles - in mediating functional interactions within the oceans. Extracellular vesicles are released by most marine microbes and are abundant in ocean waters, but our understanding of their functions remains in its infancy. As vesicles can contain diverse molecules, including active enzymes, and transport them between cells, they may work as a packaging and delivery system for goods and services traded between ecologically important microorganisms. Broader impacts of the project include providing hands-on research experiences for undergraduate and graduate students - including those from groups historically underrepresented in STEM fields - and the development of new active learning exercises to help increase knowledge about the roles microbes play in students' lives.

This project explores vesicle functions across multiple scales, combining -omics analyses, field experiments, and functional studies in cultures of diverse and ecologically important microbes to arrive at new understandings of vesicle contributions to cellular exchanges. These experiments incorporate an evolutionary perspective for exploring the range of vesicle functions and genetic mechanisms affecting their production, examining how their contents have changed in co-cultures of phytoplankton and heterotrophic bacteria following hundreds of generations of experimental laboratory evolution. Fundamental ecological questions are addressed concerning whether vesicles, and their associated functions, act as truly 'public goods' in the oceans or can instead be targeted to a subset of cells, possibly yielding 'club goods' that define interacting, cooperative networks. Collectively, this effort will generate new insights into the mechanisms marine microbes use to interact with one another, and experimentally define the functional potential and ecological impact of EV-mediated trafficking networks in the oceans.

This project is jointly funded by the Biological Oceanography Program and the Established Program to Stimulate Competitive Research (EPSCoR). This award reflects NSF's statutory mission and has been deemed worthy of support through evaluation using the Foundation's intellectual merit and broader impacts review criteria.