Enrichment of Marine Bacteriophage from Diverse

Environments and the Search for Phage-degrading

Heterotrophs

Elizabeth Ottesen

Microbial Diversity

Summer 2005

Enrichment of Marine Bacteriophage from Diverse Environments and the Search

for Phage-degrading Heterotrophs

Abstract

Phage are among the most numerous members of marine microbial communities.

The high phage population turnover rates reported in some marine environments can

represent a substantial carbon flux, and phage particles contain a high concentration of

nitrogen and phosphate. The initial purpose of this project was to evaluate the potential

of phage particles to serve as a nutrient source for marine organisms by setting up

enrichments to isolate marine organisms capable of utilizing phage as a sole source of

carbon, nitrogen, and phosphate. In preparation for this experiment, thirty different

marine bacterial isolates were challenged with phage concentrates from four different

environments. These 120 different phage-host pairs yielded 4 putative phage isolates

growing on three different strains. All successful phage enrichments utilized phage

concentrates from the environment the host was isolated from.

Introduction

In aquatic environments, viruses are the most numerous members of the microbial

community, generally present at levels of 105-108 particles per mL, and outnumbering

bacteria by 3-10 fold (Wommack and Colwell, 2000). Phage populations exhibit

dramatic spatial (Wommack et al, 1992) and temporal (Bratbak et al 1996) variation in

nature, with statistically significant changes over time intervals of 10-20 min (Bratbak et

al 1996). Phage population dynamics have been observed to mirror host populations in

both increases and decline (Muhling et al, 2005). Phage have been hypothesized to have

strong impacts on microbial community structure in the ocean through bacterial mortality

a selection (Fuhman, 1999; Weinbaur, 2004).

While the mechanisms, if not the ecology, of phage reproduction are clear, the

causes of phage ‘mortality’ are not obvious. Observed viral production and decay rates

for a variety of environments are listed in Table 1, which contains data taken from a

review by Wommack and Colwell (2000). Interestingly, viral inactivation (loss of

infectivity) and particle decay appear to occur by different mechanisms. UV damage

from sunlight is thought to be a primary cause of viral inactivation, but it was not found

to affect particle destruction rates (Wommack et al, 1996). The mechanisms of particle

loss are not clearly understood. The ability of filtration and heat treatment to decrease

rates of viral inactivation suggests a biological origin for this activity, but the source and

nature of it remains unclear. Additional factors could include adsorbtion to sediment and

colloidal organic material and protist grazing (Suttle and Chen, 1992).

The purpose of this study is to examine viral decay rates in Woods Hole marine

environments and to enrich for microbes with the ability to grow with phage particles as a

sole source of carbon, nitrogen, and phosphorus. The ability to degrade viral particles in

the surrounding environment could be of benefit to microbes both as a method of threat

removal and as a potential nutrient source. 109 particles per mL of a relatively small

virus (40kb genome) represents a carbon concentration of ~ 2 uM, as well as a rich

phosphate and nitrogen source. Assuming 50% conversion to cell mass, a bacterial cell

(2.8 x 10-13 g) would need to degrade 8.4 x 106 of these virus particles to double once, but

using the numbers listed in Table 1 and assuming steady state viral concentrations, a rich

viral community could support production of up to 2.7 x 104 cells per mL per day.

Location Production (x109 ml-1d-1)

Turnover Time (d-1)

Carbon Flux (uM d-1)

Bering/Chucki Seas 0.5-4.2 1.2-15 1-8.4 Southern California Bight Nearshore Offshore

12-230 0-2.8

.57-3.9 8.9-30

24-460 0-5.6

Santa Monica, CA 24-46 0.46-0.6 48-92 Lake Hoare, Antarctica 49 0.3 98 Table 1: Virus production and lifetime in a variety of environments. Production and turnover rate data take from a review by Wommack and Colwell (2000). Carbon flux calculated based on listed production rate and based on estimated carbon content of a phage with 40kb DNA and a 13 MDa capsid.

Methods:

All seawater isolates were grown on seawater complete (SWC) medium (.5%

tryptone, .3% yeast extract, .3% glycerol, 70% seawater). SWC plates contained 1.5%

agar, and top agar layers .6%.

Strains included a bioluminescent Vibrio fisherii sp. (strain BL2) isolated from a

10-1 dilution of water from Garbage Beach on a 3% SWC agar plate. Subsequent

transfers were grown on 1.5% agar. Additional environmental Vibrio strains were

obtained from Joanne Yeung and Erin Banning and used for host specificity assays. Also

utilized were a collection of salt-water isolates obtained from Laura Hobley. These

strains were enriched and streaked to isolation from a variety of different environments

around the Woods Hole area. Environments include a long-standing enrichment of

seawater and decomposing clams (strains 1.1-1.5), Eel Pond water (strains 2.1-2.5),

Garbage Beach water (strains 3.1-3.5), Trunk River water (strains 4.1-4.5, 4.3 not used),

Little Sippewissett empty mussel shell and water (strains 5.1-5.5), and from live mussels

and surrounding water collected at the Greater Sippewissett salt marsh (strains 6.1-6.6).

For detailed phylogenetic information regarding these strains, see Laura Hobley’s 2005

Microbial Diversity project report.

Initial enrichments for lysogenic phage were conducted on Vibrio isolate BL2

using methods detailed in the 2005 Microbial Diversity lab manual. Briefly, a phage

enrichment was set up using 50 mL water from Eel Pond, 50 mL 2x SWC medium, and

log-phase BL2 cells. This culture was then incubated for several hours at 30 deg. 1 mL

of this enrichment was then removed to a separate tube. 2 drops of chloroform were

added to promote lysis of host cells, and the mixture was then centrifuged to removed cell

debris. 100 uL of supernatant (or supernatant dilutions) was added to top agar seeded

with mid-phase log phase BL2 and plated on SWC medium. Following this step, the

100mL enrichment was allowed to stand several days at room temperature, then the

above process was repeated.

Phage were eluted from plaque-containing plates by adding 3 mL SWC media to

plates, then allowing them to stand at 4 deg C for 1-2 hr. Following this, the liquid SWC

and top agar were removed from the plates, spun in microcentrifuge tubes at 5000x for 5

min, then filtered through a .22um syringe filter to remove cells. 3mL top agar seeded

with 100uL of .3 OD cultures of host strains was spread on SWC plates and allowed to

cool, then 10 uL of filtrate and filtrate dilutions were dropped onto plates. Plates were

then tilted to allow droplets to run in a straight line across the plate, generating streaks.

Additional environmental phage isolations were conducted using Laura Hobley’s

environmental isolates. In these enrichments, 400mL water from each environment

tested was filtered through a .22um filter to remove cells. Phage were then concentrated

using a Centricon-Plus filtration device with a 30kDa molecular weight cutoff down to a

final volume of 400 uL (1000x concentration). Top agar plates for each strain were

prepared, and phage concentrates were streaked as described above. Concentrates were

diluted 10- and 100-fold in phage-free source water filtrates (retrieved from concentration

process). Plates were incubated overnight at 30 degrees, then examined for plaques.

Results

Initial enrichments for lysogenic phage capable of infecting Vibrio isolate BL2

failed. The four-hour enrichment showed no plaques. Plating additional culture medium

after extended (5 day) incubation resulted in apparent plaque formation. However, plate

eluates did not form plaques, although they were capable of inhibiting growth at high

concentrations. A possible explanation for this phenomenon is that the centrifuged but

unfiltered phage enrichments contained organisms that produced inhibitory compounds

that resulted in clearings that resembled plaques. Elution and filtration of these plaques

removed the producers but left a high enough concentration of inhibitor to reduce

somewhat the growth of cells exposed to the eluate. However, with the source organism

removed, no plaque formation could be observed.

Filtration and concentration of phage from environmental samples yielded better

results. Water from Garbage Beach, Eel Pond, Trunk River, and Little Sippewissett salt

marsh was filtered and the 30kDa – 0.22um size fraction concentrated 1000-fold. 10 uL

concentrates corresponding to 10 mL, 1 mL, and 100 uL of the source water were tested

against 30 strains, 5 from each environment and 5 each from two additional environments

(a open enrichment containing seawater and decaying clams incubated on the benchtop

for several days, and a live mussel and surrounding water from Greater Sippewissett salt

marsh). Three enrichments contained probable phages, each present in the 10 and 1 mL

streaks (Figure 1). Plaques formed on isolate 3.1, a Pseudoalteromonas strain isolated

from Garbage Beach, isolate 5.1, a marine bacillus isolated from Little Sippewissett, and

isolate 4.2, a Shewanella species isolated from Trunk River. In each case, phage were

found in water from the source environment. Isolate 4.2 exhibited what may be two

different plaque morphologies, one that was fairly turbid and one with more complete

clearing.

Discussion

Due to difficulty obtaining pure phage isolates and time limitations, I was not able

to produce sufficient phage to set up an enrichment culture. However, phage enrichments

produced some interesting data. Screening 30 isolates against 1000x concentrated 30

kDa – 0.22 um seawater particle fractions, I was able to isolate phage capable of infecting

three different strains. Each strain was sensitive only to phage present in their own

environment, despite the presence of closely related species in other environments.

Additionally, isolates with very close 16S sequences showed different sensitivities, with

isolate 4.1, virtually identical at a 16S level to 4.2, not developing plaques under identical

conditions.

The next step would be to grow and purify large quantities of these phage, and use

them to isolate microbes capable of degrading viruses by setting up enrichment cultures

using salt water basal salts media and phage at 1011 particles per mL (2mM carbon) as

sole source of C, N and P. Degradation of viruses can be monitored by enrichment

a

b

c d Figure 1: Plaque morphologies of phage isolates. Typcial plaques were shown for each isolate: Isolate 3.1 (a); Isolate 4.2 ( b, c); Isolate 5.1 (d).

growth and by quantification of plaque-forming units, and rates of degradation can be

compared to those observed in ocean water samples. Setting up these enrichments in

liquid dilution series (to avoid isolation of agar-degrading organisms) will allow

estimation of the abundance of organisms capable of degrading viral particles. If

possible, all three phage isolates should be used for enrichments. This would allow the

comparison of species isolated on each phage, and cross-comparison could be used to

evaluate ‘prey’ specificity of phage-degrading isolates.

Although enrichment of organisms on phage-containing media does not guarantee

that they are living by this mechanism in nature, these experiments should allow us to

estimate how widespread this capability is in the environment. Experiments such as

radioactive tracer studies might be preformed in order to explore the degradation of

viruses under natural conditions.

References Bratbak, G, M Heldal, TF Thingstad, P Tuomi. 1996. Dynamics of virus abundance in

coastal seawater. FEMS Microbiology Ecology. 19:263-269. Fuhrman, JA. 1999. Marine viruses and their biogeochemical and ecological effects.

Nature 399:541-548. Muhling, M, NJ Fuller, A Millard, PJ Somerfield, D Marie, WH Wilson, DJ Scanlan, AF

Post, I Joint, and NH Mann. 2005. Genetic diversity of marine Synechococcus and co-occurring cyanophage communities: evidence for viral control of phytoplankton. Environmental Microbiology. 7(4):499-508.

Suttle, CA and Feng Chen. 1992. Mechanisms and Rates of Decay of Marine Viruses in Seawater. Applied and Environmental Microbiology. 58:3721-3729.

Weinbauer, MG. 2004. Ecology of prokaryotic viruses. FEMS Microbiology Reviews. 28:127-181.

Wommack, KE, RT Hill, M Kessel, E Russek-Cohen, RR Colwell. 1992. Distribution of Viruses in the Chesapeake Bay. Applied and Environmental Microbiology. 58:2965-2970.

Wommack, KE, and RR Colwell. 2000. Virioplankton: Viruses in Aquatic Ecosystems. Microbiology and Molecular Biology Reviews. 64:69-114.

Wommack, KE, RT Hill, TA Muller, and RR Colwell. 1996. Effects of Sunlight on Bacterophage Viability and Structure. Applied and Environmental Microbiology, 62:1336-1341.