We thank Elisha Wood-Charlson, Jackie Mueller, Karen Weynberg, and Kathy Morrow for their help and constructive input on this manuscript. We also thank the crew and captain of the Hanse explorer which provided us with a perfect working environment to conduct large parts of this research. Further we would like to thank the three anonymous reviewers for their time and helpful comments to improve the manuscript. This work was funded by the NSF Dimensions of Biodiversity and PIRE awards DEB-1046413 and OISE/IIA-1243541, respectively, the Gordon and Betty Moore Foundation, Investigator Award 3781, and the CIFAR Integrated Microbial Diversity Fellowship IMB-ROHW-141679, all to FR.

1. Preparation of Instruments
<ol>
	<li>Preparation of Filtration Setup
	<ol>
		<li>Prepare a 5% hydrochloric acid (HCl) solution (0.5-0.6 M; add 250 ml 32-36% HCl to 4.75 L distilled water to make 5 L of final solution). Store this solution in a high-density polyethylene container (HDPE) for up to 2 weeks.</li>
		<li>Leave all parts that will come in contact with the sample (except the filters) for 24 hr in the 5% HCl solution to leech out potential dissolved organic carbon (DOC) contaminants. After each sampling event rinse all parts and flush lines with about 30 ml 5% HCl solution to prevent carbon contamination (e.g., from gas fumes, oil, boats, mosquito spray).</li>
		<li>Keep all sampling items wrapped in pre-combusted aluminum foil or capped until to immediately prior to use.</li>
		<li>Precombust 25 mm GF/F filters wrapped in aluminum foil in a muffle furnace at 550 &#176;C for 12 hr to volatilize off all carbon and store them in a clean, dry place until use. Use acid washed forceps and sterile, non-powdered gloves to handle the precombusted filters at all times.</li>
	</ol>
	</li>
	<li>Preparation of Viral Metagenome Sampling Materials
	<ol>
		<li>Thoroughly wash four 20 L collapsible low-density polyethylene carboys and four 20 L high density polyethylene buckets and lids with 10% bleach.</li>
		<li>Subsequently rinse all items 3x with distilled and then sample water and reseal carboys with spigot.</li>
		<li>Wash Tangential Flow Filters (TFF) after each use according to the following protocol, and clean preemptively if the filter has not been used in the past 5 days.</li>
		<li>Dissolve 50 g of NaOH pellets in 5 L of water in a wash bucket.</li>
		<li>Attach a wet TFF to the tubing and assemble according to Figure 3.</li>
		<li>Run peristaltic pump at ~30 rpm with no backpressure to move some of the NaOH wash solution through the system. Once wash flows from the return line, move transfer to wash bucket to complete the circulation of solution.</li>
		<li>Apply backpressure to the system by placing a clamp on the return line downstream of the TFF. This will force wash solution out the filtrate line. Excessive time (5 min) to flood the outside of the TFF and produce filtrate is indicative of a degraded TFF.</li>
		<li>Run the system until all of the NaOH solution is in the lines. Then remove backpressure, run solution out of the system and discard.</li>
		<li>Rinse the TFF by running water under backpressure until the pH of the water flowing from the return and filtrate lines is pH 7-8.</li>
		<li>Remove tubing from the peristaltic pump and TFF. Reseal TFF for storage. Ensure that the TFF remains wet until the next use.</li>
	</ol>
	</li>
</ol>
2. Sample Collection
NOTE: During transport all collected samples should be stored cool (if possible 4 &#176;C) and not exposed to direct sunlight until further processing.
<ol>
	<li>Sampling for Carbon, Nutrients, Microscopy, Flow Cytometry, and Microbial Metagenomics
	<ol>
		<li>Take two Hatay Niskin units per sampling event (resulting in 4 L sample water). Open the sampling vessels just before submerging (otherwise trapped air will prevent descending).</li>
		<li>At the respective site, flush the inside of the sampling vessel with the sample water by opening both ends and moving the cylinder along the polar axis through the water.</li>
		<li>Take the sample and carefully close the sampling container. Make sure the sampling site is upstream from boat and divers to avoid contamination.</li>
	</ol>
	</li>
	<li>Sampling for Viral Metagenomics
	<ol>
		<li>On the target site mount bilge pump on 20 L carboy.</li>
		<li>Fill up carboy aiming bilge pump inlet at the each targeted area and seal carboy with the corresponding spigot.</li>
		<li>Collect 4 carboys of unfiltered seawater (all together 80 L) from each sampling site.</li>
	</ol>
	</li>
</ol>
3. Sample preparation
NOTE: Process the samples in the order presented here, starting with DOC to minimize the chance of contamination. Use powder free gloves for the entire procedure.
<ol>
	<li>Dissolved Organic Carbon (DOC)
	<ol>
		<li>Mount one of the two aliquot Hatay Niskin units in respective slot of the filtration set. Connect the outlet tubing which leads to the respective filter holder. Connect the pressurized air (0.2 bar) tubing to the Hatay Niskin unit (See Figure 2).</li>
		<li>Flush all lines with 100 ml of sample water before inserting filters in the filter holders. Place 25 mm pre-combusted GF/F filter in filter holder using the acid washed forceps. Rinse each DOC bottle and cap 3 times with about 20 ml filtered sampling water.</li>
		<li>Fill bottle with about 40 ml (~2/3 full) of sampling water. Collect at least duplicates of DOC samples to have a backup in case of potential contamination or loss during transport.</li>
		<li>Freeze sampling bottles standing upright at -20 &#176;C until analysis.</li>
	</ol>
	</li>
	<li>Particulate Organic Matter (POM)
	<ol>
		<li>After DOC samples have been collected continue filtering until a total of 500 ml have passed the GF/F filter, including what passed through while collecting DOC samples.</li>
		<li>Unscrew inline filter holder.</li>
		<li>Remove the filter for POM analysis using forceps and place filter inside a square of pre-combusted aluminum foil, fold top-side turned in on itself, and wrap.</li>
		<li>Freeze filters at -20 &#176;C for storage until subjected to elemental and isotopic analysis.</li>
	</ol>
	</li>
	<li>Inorganic Nutrients
	<ol>
		<li>Place a 0.2 &#181;m Track-Etched filter into each filter holder. Re-attach the filter cassettes.</li>
		<li>Rinse each 20 ml plastic scintillation bottle three times with sampling water. Fill each bottle to the shoulder (~18 ml).</li>
		<li>Freeze at -20 &#176;C until later analysis.</li>
	</ol>
	</li>
	<li>Microscopy (SYBR Gold and DAPI)
	<ol>
		<li>Take off filter holder.</li>
		<li>Collect 1 ml of water in each of two microcentrifuge tubes.</li>
		<li>Add 66 &#181;l 32% paraformaldehyde to one of the aliquots (for later SYBR Gold staining). Add 12 &#181;l 25% glutaraldehyde to the corresponding other (for later DAPI staining).</li>
		<li>Gently mix, and allow samples to &#8220;fix&#8221; for at least 15 min at RT in the dark.</li>
	</ol>
	</li>
	<li>Flow Cytometry
	<ol>
		<li>Place an 8.0 &#181;m polycarbonate filter in one of the filter holders to exclude debris and large eukaryotic cells.</li>
		<li>Pass sampling water through to fill 2 cryovials from each sampling site with 1 ml sample.</li>
		<li>Add 5 &#181;l of 25% glutaraldehyde to each cryovial (final concentration = 0.125%).</li>
		<li>Invert vials to mix.</li>
		<li>Allow the sample to fix for 15-30 min at RT. Do not exceed 30 min.</li>
		<li>Flash freeze samples in liquid nitrogen.</li>
		<li>Store samples at -80 &#176;C or in liquid nitrogen dry shipper until analysis on a flow cytometer.</li>
	</ol>
	</li>
	<li>Microbial Metagenomic Sample
	<ol>
		<li>Remove in line filter holders.</li>
		<li>Directly mount a 0.22 &#181;m cylindrical filter onto the respective line.</li>
		<li>Filter remaining sample water of both Hatay Niskin units from each site (totaling 3-4 L per filter) through one filter.</li>
		<li>After the filtration push the remaining water out of each filter by using a clean 10 ml syringe filled with air.</li>
		<li>Place filter back into original packaging and seal the package with lab tape.</li>
		<li>Store individually packed filters at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Viral Metagenomic Sample
	<ol>
		<li>Transfer viral metagenome samples immediately into the washed buckets to ensure no sample is lost or contaminated by the environment.</li>
		<li>Pre-filter using large pore size nylon mesh (25-100 &#181;m) to remove debris and cellular material prior to concentration19.</li>
		<li>Set up the TFF as shown in Figure 3, placing the delivery line in a sample bucket, and leaving the return and filtrate lines running into a sink.</li>
		<li>Turn on peristaltic pump and flush lines with 1-2 L of sample water.</li>
		<li>Place return line in the sample bucket to complete cycle, add 0.7 bar of backpressure.</li>
		<li>While concentrating the seawater, top up sample reservoir as the level drops. When water level drops below the intake line in the bucket, transfer concentrate into a bleach-washed, triple rinsed tripour beaker and continue concentrating.</li>
		<li>If reservoir beaker is empty, remove backpressure, increase the pump rate and push the entire sample through the lines, recovering it in the tripour.</li>
		<li>Pass the concentrate through 0.45 &#181;m cylindrical filters to remove most Bacteria while not discriminating against any viral lineages.</li>
		<li>Change 0.45 &#181;m cylindrical filters after every 150 ml. Collect the 0.45 &#181;m-filtered viral concentrate in 50 ml tubes.</li>
		<li>Add 250 &#181;l of chloroform to each 50 ml aliquot of filtered viral concentrate to eliminate residual Bacteria. Invert to mix, and store, upright at 4 &#176;C until subsequent processing.</li>
		<li>Dry the 0.45 &#181;m filters and store them as described in Steps 3.6.5 and 3.6.6.</li>
	</ol>
	</li>
</ol>
4. Processing of Microscopy Samples
NOTE: Rinse filter towers with 10% bleach followed by 95% ethanol to prevent potential stain or biological residues between runs. Process microscope samples within 1 hr and avoid exposing stains and stained filters to light if possible.
<ol>
	<li>Mount
	<ol>
		<li>Add 100 &#181;l of 10% ascorbic acid to 4.9 ml of 1x PBS, and mix thoroughly.</li>
		<li>Add 5 ml of 100% glycerol, and mix thoroughly.</li>
		<li>Filter mount using a 0.02 &#181;m alumina matrix disposable syringe filter, aliquot and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>SYBR Gold Staining
	<ol>
		<li>Pipette a 500 &#181;l aliquot of each sample fixed with 2% final concentration paraformaldehyde in a microcentrifuge tube.</li>
		<li>Add 0.5 &#181;l of 1,000x SYBR Gold solution to each of the aliquots.</li>
		<li>Mix sample gently and put it in the dark for 10 min.</li>
		<li>Place a 0.02 &#181;m alumina matrix filter with annular polypropylene support ring on each filter stand of the vacuum pump system and attach the filter towers (Figure 4). Make sure the 0.02 &#181;m filter is placed on the filter stand plastic-side up, as these filters have a dual porosity.</li>
		<li>With the vacuum pump turned off, add the 500 &#181;l of SYBR stained sample, along with 2 ml of 0.02 &#181;m filtered virus free water to ensure the sample is spread evenly across the filter.</li>
		<li>Turn on the vacuum pump to create a negative pressure of no more than 1 bar to avoid damaging viruses and microbes being filtered.</li>
		<li>Allow the entire sample to go through.</li>
		<li>Using forceps (not the same pair as used for DOC), carefully remove each filter from filtering tower, and pass over a delicate task wipe to ensure the filter is dry.</li>
		<li>Pipette 10 &#181;l of mount on a microscope slide and place the dry filter containing the stained sample on the drop of mount.</li>
		<li>Add 10 &#181;l mount to the top of each filter before adding a coverslip.</li>
		<li>Gently press coverslip down. Try to eliminate air-bubbles if possible and avoid sideways motion of the coverslip.</li>
		<li>Place slide in a slidebox and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>DAPI Staining
	<ol>
		<li>Place a 0.2 &#181;m alumina matrix filter with annular polypropylene support ring on each filter stand and attach the filter towers.</li>
		<li>With the vacuum pump turned off, add 2 ml of 0.02 &#181;m filtered virus free water to each tower used.</li>
		<li>Pipette 1 ml of the glutaraldehyde-fixed sample into each tower, resulting in a total volume of 3 ml in each filter tower.</li>
		<li>Turn on vacuum pump to create a negative pressure of no more than 1 bar.</li>
		<li>Filter entire sample onto the 0.2 &#181;m alumina matrix filter.</li>
		<li>Carefully remove filter from filtering tower using forceps (not the same pair as used for DOC).</li>
		<li>Place the filter on a 100 &#181;l drop DAPI solution [25 &#181;g/ml] in a Petri dish keeping the filter right side up and let the sample stain for 15 min. Avoid light during the staining process.</li>
		<li>After the 15 min, remove the filter from the stain, pass it over a delicate task wipe to dry and transfer to a 100 &#181;l drop of 0.02 &#181;m filtered virus free water, again keeping the filter right side up, to wash off remaining DAPI stain for 1 min. Repeat this wash once.</li>
		<li>Mount filter on microscope slide the identical way as described for the SYBR Gold samples.</li>
	</ol>
	</li>
</ol>
5. DNA Extraction
<ol>
	<li>Microbial Metagenomic DNA
	<ol>
		<li>Thaw the cylindrical filters for at least 20 min at RT.</li>
		<li>Remove any remaining seawater in the filter using a 10 ml syringe. Cap the bottom end of the cylindrical filter using a bleached and autoclaved cap.</li>
		<li>For each filter, mix 360 &#956;l T1 &#8220;pre-lysis&#8221; buffer and 50 &#956;l Proteinase K, and then pipette the mixture into the filter. Cap the open end.</li>
		<li>Incubate the lysis reaction in a rotating oven O/N (or at least 2 hr) at 55 &#176;C.</li>
		<li>Remove one of the cap and add 200 &#181;l B3 &#8220;lysis&#8221; buffer into the filter to the pre-lyse sample. Re-cap filter and incubate the mixture in a rotating oven for 10-20 min at 70 &#176;C.</li>
		<li>Extract the entire volume from the cylindrical filter using a 3 ml syringe by sucking the liquid out with the filter upside down.</li>
		<li>Expel the lysed sample into a new microfuge tube, and add 200 &#956;l of 100% ethanol.</li>
		<li>Mix well, load the entire sample onto the spin column, and proceed as instructed by the manufacturer&#39;s protocol.</li>
		<li>Quantify DNA using a fluorescent-based assay.</li>
	</ol>
	</li>
	<li>Marine Viral Metagenomics
	<ol>
		<li>Purifying and Concentrating Phage Particles (modified from Thurber et al.19)
		<ol>
			<li>Dissolve CsCl in seawater to prepare solutions of 1.7 g/ml, 1.5 g/ml, 1.35 g/ml, 1.2 g/ml (calibrated by weighing out 1 ml of solution). Filter each fraction with 0.02 &#181;m filter prior to use. 
			NOTE: Ensure that the density of each fraction is accurate to three significant figures before using the solutions and filter all solutions with a 0.02 &#181;m filter before use. Use seawater or seawater saturated with CsCl to lower or increase density, respectively.</li>
			<li>Make a microscope slide from 1 ml of combined reagents as described in section 4.2 to ensure that all reagents are devoid of viruses before proceeding.</li>
			<li>Add 9 g CsCl to 45 ml of viral TFF concentrate to a final density of 1.12 g/ml. Refrigerate while setting up the CsCl step gradient.</li>
			<li>Starting with the 1.7 g/ml solution successively add 1 ml of each sequentially less dense fraction slowly into each tube using serological pipettes. Mark the level of each individual meniscus and ensure that the pycnoclines between fractions are not disturbed. For more detail see the companion paper (Lim et al. 2014).</li>
			<li>With a serological pipette load 7.5 ml of sample into each of 6 tubes and centrifuge at ~83,000 x g, 4 &#176;C for 2 hr.</li>
			<li>Without disrupting the density gradients unload the rotor and pierce the tube just below the previously marked 1.5 g/ml density layer (Figure 5) with an 18 G needle on a 3 ml syringe. Draw off 1.5 ml of purified VLPs and transfer to a new microcentrifuge tube. Repeat for all samples.</li>
			<li>Add 0.2 volume of chloroform to the purified VLPs, mix vigorously, incubate at RT for 10 min.</li>
			<li>Add 150 &#181;l of 10x DNAse buffer to each microcentrifuge tube.</li>
			<li>Spin at 16,000 x g for 5 min, and collect the aqueous phase. If the chloroform does not aggregate in the bottom of the microcentrifuge tube, add more DNAse buffer, mix and repeat.</li>
			<li>Add DNase I (final concentration = 2.5 unit/&#181;l) and incubate at 37 &#176;C for 2 hr, then inactivate DNase activity by incubating at 65 &#176;C for 15 min.</li>
		</ol>
		</li>
		<li>DNA Extraction from Viral-like Particles (VLPs)
		<ol>
			<li>Evenly pool the purified VLPs from each sample into two cleaned and autoclaved Oak Ridge tubes.</li>
			<li>Add 0.1 volume 2 M Tris-HCl (pH 8.5) with 0.2 M EDTA, 0.01 volume 0.5 M EDTA, 1 volume of formamide, 10 &#956;l glycogen (10 mg/ml). Mix well and incubate at RT for 30 min.</li>
			<li>Referring to new volume, add 2 volumes RT 100% ethanol. Mix and incubate at 4 &#176;C for more than 30 min.</li>
			<li>Pellet DNA by spinning the Oak Ridge tubes at 17,200 x g for 60 min, at 4 &#176;C.</li>
			<li>Remove the supernatant. Wash the DNA pellet twice with ice-cold 70% ethanol using serological pipettes.</li>
			<li>Remove all ethanol (re-spinning briefly if necessary), cover, and allow pellet to air-dry at RT for &#62;15 min.</li>
			<li>Resuspend the DNA pellet for &#62;15 min in 567 &#956;l of 1X TE buffer (pH 8.0). Transfer the 567 &#956;l of resuspended DNA into a new 1.5 ml microcentrifuge tube.</li>
			<li>Add 30 &#956;l of 10% SDS (pre-warm at 65 &#176;C prior to use) and 3 &#956;l of proteinase K (20 &#956;g/ml), mix thoroughly and incubate for 1 hr at 56 &#176;C.</li>
			<li>Add 100 &#956;l of 5 M NaCl and mix thoroughly. Add 80 &#956;l of pre-warmed CTAB NaCl solution, vortex, and incubate for 10 min at 65 &#176;C.</li>
			<li>Add 0.2 parts of volume of chloroform, vortex, and spin at 16,000 x g for 2 min. Transfer supernatant to a new 1.5 ml microcentrifuge tube.</li>
			<li>Add equal volume of phenol chloroform isoamyl 25:24:1 solution, vortex to mix, and spin at 16,000 x g for 2 min. Transfer the supernatant to a new 1.5 ml microcentrifuge tube.</li>
			<li>Add equal volume of chloroform, vortex, and spin at 16,000 x g for 2 min. Transfer the supernatant to a new 1.5 ml microcentrifuge tube.</li>
			<li>Add 0.7 volume of isopropanol to the supernatant fraction and incubate at -20 &#176;C for at least 30 min to precipitate DNA.</li>
			<li>Pellet the DNA by spinning at 16,000 x g for 20 min at 4 &#176;C. Pipette off the supernatant and wash the pellet twice with 500 &#181;l of ice-cold 70% ethanol.</li>
			<li>Spin at 16,000 x g and remove the remaining ethanol from the tube. Air-dry the pellet for 15 min.</li>
			<li>Resuspend DNA pellet with 50 &#956;l of elution buffer (5 mM Tris, pH 7.5) or Tris-EDTA pH 8.0 for at least 5 min at RT.</li>
			<li>Quantify the DNA using high-sensitivity fluorescence-based assay.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The methods presented here will provide a tool to generate a comprehensive assessment of viral and microbial dynamics in aquatic ecosystems. They are targeted to quantify the standing stock of organic and inorganic resources available to microbial communities and to characterize the makeup of the viral and microbial community present. Next to quantifying viral abundance and microbial abundance and biomass, genomic sequence information reveals their community structure and function. All methods have been developed or modified to allow for application in remote field locations. However the lack of controlled laboratory settings inherently creates some potential for errors. Here we discuss some caveats associated with each of the parameters assessed. Next to potential for contamination during sample handling some of the parameters also yield potential for error in the analytical process.
Dissolved Organic Carbon
Carbon contamination can occur during the sampling procedures (sampling downstream, or in the vicinity of a boat or diver), during the sample preparation (contamination of equipment or inadvertent handling), and even during storage of the sample (other samples in freezer containing organic solvents/volatiles). To avoid the various sources of contamination conduct as few handling steps as possible, as each sample relocation poses an additional source of contamination. The sample should not get in contact with any item not acid washed or combusted. Finally, designating a storage freezer exclusively for samples not containing volatile organic substances will ensure the integrity of the samples during longer storage periods. If samples cannot be kept frozen during an extended travel period acidification of DOC samples with concentrated HCl (as described38-41) may be an applicable alternative. To verify the measurement accuracy DOC consensus reference materials should be used regularly during the DOC measurement runs. Especially deep sea water (&#62;2,000 m) references are valuable in assessing the performance of the analytical machine as it is very stable in its DOC content42.
Particulate Organic Matter
In addition to avoiding organic carbon contamination, POM sampling requires special attention to ensure the accuracy of the filtered volume. If the targeted volume of 500 ml should deviate for any potential reason this needs to be noted to relate the amount of POM captured on the filter to the filtrate of which it was derived.
Inorganic Nutrients
As with organic carbon sampling, attention must be paid to possible nutrient sample contamination generated by various sources like boats or wastewater discharges12. Filters used for organic carbon sampling with a nominal pore sizes of 0.8 &#181;m, may not exclude all microbial biomass and should therefore be changed to 0.2 &#181;m filters for this filtration step. Further, detection limits pose a problem with nutrient samples; especially in the oligotrophic surface water around many coral reef locations (e.g., Cotner&#160;et al.43). Detection limits are roughly around 0.1, 0.2, and 0.1 &#956;mol/L&#160;for ammonium, nitrate and nitrite, and ortho-phosphate respectively (see 36,12,37)
Microscopy
Beside variations in the concentration of samples image analysis from microscopy slides further yields potential for errors. For instance, images may be out of focus in some areas of the field of view, and in focus in others. As a high-purity alumina matrix filter is a very rigid filter type, any debris under the filter may cause the whole filter to sit on an angle to the slide. As a result, images can be consistently out of focus in various parts of the field of view for a given filter, regardless of microscope alignment. Remounting filters, ensuring the underside of the filter is clean, can ameliorate this if it is observed. Also, in some cases there may be two focal planes in which viruses and microbes can be found. This is the result of stained objects becoming detached from the filter upon mounting, and floating up to the underside of the cover slip which can make counts unreliable. Therefore it is always recommended to check the quality of the samples during the process.
Flow Cytometry
As with the microscopy samples, there may be naturally occurring differences between flow cytometry samples which require adjustments of the analytical process. For instance, depending on the sensitivity of the instrument, dimly fluorescing Prochlorococcus cells may be below the noise level&#160;and will not be quantified. Beads serve as an internal sample control (e.g., to confirm that the instrument&#8217;s response to fluorescent signals is consistent from day to day (and remains stable during the run itself). If added to the sample at known concentrations, they can also serve as an internal control to verify the sample volume run. However, it is recommended to always run an aliquot of a previously frozen &#8220;standard&#8221; seawater sample that is thawed and stained along with the samples of interest to provide an additional biological control for sample handling between 96-well plate runs. Depending on the location, seawater samples may look very different from each other. Sorting &#8220;representative&#8221; populations and then viewing under an epifluorescent microscope (EM) can be a good way to quickly verify phytoplankton populations as either Synechococcussp. or photosynthetic eukaryotes. Prochlorococcus cells are typically not visible under EM because they fade too fast. In addition to chlorophyll a, Synechococcussp. also contain the photopigment phycoeurythrin (PE), which emits light at shorter wavelengths (540-630 nm) than chlorophyll A (660-700 nm). When Synechococcus cells are excited with blue/green light (470-490 nm), they will appear golden if viewed under an emission filter that removes all wavelengths of light below 510 nm. By comparison, the eukaryotic fraction will look red, when excited with the same wavelength of light.
Viral Metagenomics
It is important to note that the process of VLP concentration and storage influences the community recovered. Viral particle loss can occur at every step in the process, and thus,&#160;the selection of viral purification and enrichment protocols can ultimately affect the observed taxonomic composition and diversity of the resulting viral metagenomes44,45,18. Concentration of samples can be done using TFF19 or chemical-based flocculation20. In the chemical-based approach, positively charged iron ions bind the naturally negatively charged viral particles forming large (&#62;8 &#181;m) iron-viral complexes that flocculate out of solution and can be recovered using 8 &#181;m filters. Subsequently the iron is chelated off the viral particles and re-dissolved using magnesium, ascorbic acid, and EDTA, leaving concentrated viral particles for nucleic acid extraction20. After comparing these two current methods, we concluded that the iron chloride method is less time consuming than TFF viral concentration, but may yield some caveats. We found that dissociation and dissolution of the iron-viral requires a two-fold more concentrated magnesium-ascorbic acid-EDTA solution than reported20 in order for dissolution to proceed before the ascorbic acid degrades. Aside from this issue, the protocol purifies viral particles by size-fractionation alone, removing microbial contaminants using 0.2 &#181;m filtration. Large viruses do not pass through the filter (e.g., Yang&#160;et al.46) and will thus not be detected in the metagenome, while some Bacteria do (McDole, unpublished data). This results in bacterial contamination while discriminating against large viruses.
That specific issue however is also relevant for the removal of microbes in connection with TFF concentration. As the chloroform treatment has been shown to remove chloroform-sensitive viruses with external lipid membranes47 some studies suggest that 0.22 &#181;m filtration can be used to remove the majority of bacteria. It should also be noted that the presented method primarily targets bacteriophage, the most abundant carrier of genetic material in marine environments. Because phage are generally thought not to commonly contain lipids 48&#160;they are more resistant to chloroform treatment. To our knowledge a &#8220;catch-all&#8221; method does not exist to date. The method presented here is adapted from our field experience over the last 10 years in various marine environments spanning tropical to arctic systems.
Microbial Metagenomics
The construction of metagenomic libraries for microbes (i.e., bacteria and archaea) is more straightforward than viral metagenomes as it is easier to generate the minimal DNA concentrations required for library preparation. Linker amplification shotgun libraries (LASLs)17,49,50 and whole genome amplification based on multiple displacement amplification (MDA) are the two most commonly used methods to generate sufficient DNA for sequencing. The use of MDA to obtain higher DNA concentrations in microbial metagenomes is sometimes necessary. This however can cause artifacts in the sequence data, such as the overamplification of dinoflagellate minicircles. MDA methods are known to preferentially amplify single-stranded and circular DNA, resulting in non-quantitative results51,52. The optimized LASLs approach50 may thus serves as a better alternative in amplifying both microbial and viral metagenomic DNA for sequencing. It is important to note that the LASLs approach has multiple steps, requires sophisticated equipment, and is limited to dsDNA templates. Further, the tissue DNA extraction kit has been shown to extract DNA from both Gram positive and Gram negative phyla, but it has not been verified to effectively lyse recalcitrant microbial taxa or their associated structures (e.g., certain archaea, endospores, or fungal spores). Therefore a bead beating step may be necessary to obtain DNA from certain microbes.
These comprehensive assessments will result in a better understanding of viral and microbial ecosystem functioning. Although few studies have undertaken the effort to assess all proposed parameters, many have been investigating selected ones. Nelson et al.53 suggested a connection between specific reef habitats and depletions in both DOC and bacterioplankton concentrations relative to offshore waters. These concentration changes were accompanied by distinct differentiations of bacterioplankton communities. Dinsdale et al.5 showed that increased anthropogenic impact was accompanied by 10x&#160;higher abundances of microbial cells and virus-like particles. Microbial communities in marine waters surrounding inhabited islands also had larger fractions of heterotrophs and potential pathogens5. Finally McDole et al.4 showed, by introducing the microbialization score, that human activities are shifting energy to the microbes, at the expense of the macrobes. These studies, focused on specific microbial and water chemistry parameters, provide novel insights into the biochemical structure of marine ecosystems. Combining traditional data of ecosystem monitoring efforts - like temperature and pH values, fish biomass and diversity, benthic cover, or exposure to human impact - with water chemistry and microbial assessments, will likely result in more sensitive environmental monitoring efforts, which may lead to more specifically targeted conservation efforts.

Marine ecosystems are subjected to a wide range of perturbations that result in changes from nutrient availability to the biomass of apex predators. Over the last decade, multiple studies have demonstrated the importance of the microbial communities in marine ecosystems1-4. It is evident that changes in the bacterial and viral community are closely associated with the overall degradation of marine environments5. These changes may be facilitated by altered geochemistry in the water column such as oxygen availability or carbon remineralization 6,7. As a result of the interdependency between the macroorganisms, water chemistry and microbial activity feedback loops may accelerate the rate of ecosystem degradation8.
In the 1970s and 80s multiple attempts were made to unravel biogeochemical cycles in marine ecosystems9-11. One of the main challenges for these early studies was the lack of standardized approaches to measure the assessed biogeochemical parameters. Though there were protocols available, primarily on seawater nutrients12,13, the assessment of carbon dynamics and the microbial community structure were limited by the tools and methods available. With the JGOFS EqPac methods comparison, Sharp et al.14 suggested for the first time a reliable and standardized protocol for dissolved organic carbon (DOC) concentration assessments. By the early 2000s the analytical challenges to determine resources available to the microbial community had thus been largely resolved and approaches to characterize microbial communities in controlled culture environments had been established (see DeLong15). The traditional sequencing methods at that time largely relied on cultivated clonal cultures. Early environmental gene sequencing of natural samples revealed, however, that the majority of microbial biodiversity had been missed by cultivation-based methods16. In 2002, Breitbart et al.17 used shotgun sequencing for the first time to describe the viral community in environmental seawater samples establishing a method to sequence the entire genome of uncultured marine viral communities.
Within the existing microbial assessments, viruses still remain particularly under-studied, as most are difficult to culture and they lack a universally conserved marker such as the 16S ribosomal RNA (rRNA) genes typically used for assessing diversity and community profiling. The metagenomic sequencing approach provides an alternative to culture-dependent and gene-targeted methods for analyzing complex viral communities. While the first viral metagenomes generated from seawater were sequenced using Sanger sequencing17, the development of next-generation sequencing technologies and improved molecular techniques has led to a rapid increase in metagenomic studies18. The current laboratory workflow for viral metagenomics involves the separation, enrichment, and/or concentration of viral particles, followed by nucleic acid (DNA or RNA) extraction, library preparation, and sequencing19-21.
To further the existing knowledge on viral, microbial and biochemical functioning in aquatic ecosystems, comparing data sets in various systems around the world is essential. However, the established protocols have largely been developed for near-shore environments with readily accessible laboratory facilities. Thus, the lack of standardized field methods hinders these cross-ecosystem comparisons. Here we introduce thoroughly tested and well standardized adaptations from state of the art research protocols for use in remote field locations. These methods have been proven successful in multiple studies spanning the last decade5,22&#160;and are currently used by various marine laboratories as well as on the NOAA Reef Assessment and Monitoring Program (RAMP) throughout the Pacific Ocean4. The sampling protocols include water chemistry parameters (inorganic and organic carbon concentrations, inorganic nutrient concentration), viral and microbial abundance (direct bacterial counts, direct viral counts, and flow cytometry to assess autotroph vs heterotroph ratios), and viral and microbial community structure and metabolic potential through metagenomic analysis (for an overview see Figure 1). The results obtained from the methods described&#160;here are required in order to elucidate key biogeochemical background parameters needed to comprehensively characterize aquatic ecosystems.

<table border="0" cellpadding="0" cellspacing="0" width="1307">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:616px;">Name of Material/ Equipment</td>
			<td style="width:217px;">Company</td>
			<td style="width:149px;">Catalog Number</td>
			<td style="width:325px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Niskin collection and processing units (Hatay Niskins)</td>
			<td style="width:217px;"></td>
			<td style="width:149px;"></td>
			<td>Figure 1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Filtration setup&#160;</td>
			<td style="width:217px;"></td>
			<td style="width:149px;"></td>
			<td>Figure 1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">32 &#8211; 36 % Hydrochloric acid (HCl)</td>
			<td>Fisher Sci</td>
			<td>A508-212</td>
			<td></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:18px;">High-density polyethylene (HDPE) container</td>
			<td style="width:217px;"></td>
			<td style="width:149px;"></td>
			<td>e.g., 5 Gallon HDPE UN Certified Pail with Snap-On Lid</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Combustion oven (550&#176; C)</td>
			<td style="width:217px;"></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">60 ml HDPE vials&#160;</td>
			<td>Fisher Sci</td>
			<td style="width:149px;">02-896-2B</td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">25 mm GF/F filter</td>
			<td>Fisher Sci</td>
			<td>09-874-64</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Forceps&#160;</td>
			<td>Fisher Sci</td>
			<td>XX62-000-06</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sterile, non-powdered gloves</td>
			<td>Fisher Sci</td>
			<td style="width:149px;">19-170-010B</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bilge pump&#160;</td>
			<td></td>
			<td style="width:149px;"></td>
			<td>e.g., Thirsty Mate 124PF</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">20 l collapsible low-density polyethylene carboy&#160;</td>
			<td></td>
			<td style="width:149px;"></td>
			<td>e.g., Reliance Fold A Carrier II</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tangential flow filter (TFF) ultrafiltration hollow fiber cartridges</td>
			<td>Amersham</td>
			<td>CFP-2-E-9A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Platinum-cured silicon tubing for TFF</td>
			<td>Cole Parmer</td>
			<td>96440-82</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Hose clamps</td>
			<td>Cole Parmer</td>
			<td>P-68007-23</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Peristaltic pump</td>
			<td>Cole Parmer</td>
			<td>EW-77410-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium hydroxide (NaOH) solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>21162</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.2 &#181;m polycarbonate Track-Etched Membrane filter</td>
			<td>Fisher Sci</td>
			<td>09-300-61</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">8.0 &#181;m polycarbonate Track-Etched Membrane filter</td>
			<td>Fisher Sci</td>
			<td>09-300-57</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.22 &#181;m polyvinylidene difluoride cylindrical filter</td>
			<td>Fisher Sci</td>
			<td>SVGP01050</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.45 &#181;m polyvinylidene difluoride cylindrical filter</td>
			<td>Fisher Sci</td>
			<td>SVHV010RS</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Synthetic nylon mesh</td>
			<td>Sefar</td>
			<td>03-125-37</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">20 ml plastic scintillation bottle&#160;</td>
			<td>Fisher Sci</td>
			<td>03-341-72C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Clean bucket (10 - 20 L)</td>
			<td style="width:217px;"></td>
			<td style="width:149px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">32 % paraformaldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>15714</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">25 % glutaraldehyde</td>
			<td style="width:217px;">Electron Microscopy Science</td>
			<td>16220</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Liquid Nitrogen</td>
			<td style="width:217px;"></td>
			<td style="width:149px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.02 &#181;m high-purity alumina matrix filter with annular polypropylene support ring&#160;</td>
			<td>Fisher Sci</td>
			<td>6809-6002</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.2 &#181;m&#160; high-purity alumina matrix filter with annular polypropylene support ring&#160;</td>
			<td>Fisher Sci</td>
			<td>6809-6022</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">10,000 X SYBR Gold nucleic acid stain&#160;</td>
			<td>Invitrogen</td>
			<td>S11494</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">DAPI solution 25 &#181;g ml-1</td>
			<td>Invitrogen</td>
			<td>D1306</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Molecular grade water</td>
			<td>sigma aldrich</td>
			<td>W-4502</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ascorbic Acid</td>
			<td>Fisher Sci</td>
			<td>BP351-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phosphate buffered saline (PBS)</td>
			<td>Fisher Sci</td>
			<td style="width:149px;">BP399-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glycerol</td>
			<td>Fisher Sci</td>
			<td>G31-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Microscope slide</td>
			<td>Fisher Sci</td>
			<td>12-550-123</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Coverslip</td>
			<td>Fisher Sci</td>
			<td>12-548-5M</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Delicate task wipe&#160;</td>
			<td>Fisher Sci</td>
			<td>06-666A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Slide Station with vacuum pump</td>
			<td style="width:217px;"></td>
			<td style="width:149px;"></td>
			<td>Figure 3</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:616px;">DNA extraction kit</td>
			<td style="width:217px;">Macherey-Nagel</td>
			<td>740952.1</td>
			<td>Nucleospin tissue kit</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">Proteinase K (20 mg ml-1)</td>
			<td style="width:217px;">Lifetechnologies</td>
			<td>AM2546</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">200-proof 100% Ethanol</td>
			<td>Electron Microscopy Sciences</td>
			<td>15058</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Red cap: Male Luer with Lock Ring x Female Luer Coupler&#160;</td>
			<td>Cole Parmer</td>
			<td>EW-45503-88</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cesium chloride (CsCl)</td>
			<td>Fisher Sci</td>
			<td>bp1595-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">60 ml syringe</td>
			<td>Fisher Sci</td>
			<td>13-6898</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.02 &#181;m alumina matrix disposable syringe filter</td>
			<td>Fisher Sci</td>
			<td>O992613</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:616px;">Ultra-clear centrifuge tubes</td>
			<td style="width:217px;">Beckman Coulter</td>
			<td>344059</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ultra - swinging bucket rotor</td>
			<td style="width:217px;">Beckman Coulter</td>
			<td style="width:149px;">333790</td>
			<td>SW41 Ti Rotor&#160;</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">DNase I (100 u &#181;l-1)</td>
			<td>Calbiochem</td>
			<td>260913</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.5 M EDTA</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">BP2482-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Formamide</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">BP228-100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glycogen</td>
			<td style="width:217px;">sigma aldrich</td>
			<td style="width:149px;">G-1767</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">100 % Ethanol (200-proof)</td>
			<td style="width:217px;">Electron Microscopy Science</td>
			<td style="width:149px;">15058</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1 X Tris-EDTA (TE) pH 8.0</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">BP2473-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Dodecyl Sulfate (SDS)</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">02674-25</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">20 &#956;g ml-1 Proteinase K</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">BP1700-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Chloroform</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">BP1145-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phenol:Chloroform:Isoamyl alcohol 25:24:1</td>
			<td style="width:217px;">sigma aldrich</td>
			<td style="width:149px;">p3803</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isopropanol</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">BP2618-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:616px;">50ml Oak Ridge Tube&#160;</td>
			<td style="width:217px;">Fisher Sci</td>
			<td style="width:149px;">05-562-16B</td>
			<td></td>
		</tr>
	</tbody>
</table>

unraveling the unseen players in the ocean - a field guide to water chemistry and marine microbiology

Here we introduce a series of thoroughly tested and well standardized research protocols adapted for use in remote marine environments. The sampling protocols include the assessment of resources available to the microbial community (dissolved organic carbon, particulate organic matter, inorganic nutrients), and a comprehensive description of the viral and bacterial communities (via direct viral and microbial counts, enumeration of autofluorescent microbes, and construction of viral and microbial metagenomes). We use a combination of methods, which represent a dispersed field of scientific disciplines comprising already established protocols and some of the most recent techniques developed. Especially metagenomic sequencing techniques used for viral and bacterial community characterization, have been established only in recent years, and are thus still subjected to constant improvement. This has led to a variety of sampling and sample processing procedures currently in use. The set of methods presented here provides an up to date approach to collect and process environmental samples. Parameters addressed with these protocols yield the minimum on information essential to characterize and understand the underlying mechanisms of viral and microbial community dynamics. It gives easy to follow guidelines to conduct comprehensive surveys and discusses critical steps and potential caveats pertinent to each technique.

Microscopy
Microscopy samples can and should be analyzed immediately to ensure they are of the desired quality. To measure abundance and size distribution of the bacterial community, DAPI slides will be examined by epifluorescence microscopy (excitation/emission: 358/461 nm, see McDole et al. 2012) (Figure 6A). Cell counts and dimensions can be collected using imaging software (e.g., ImagePro or ImageJ). From length and width measurements cell volumes (V) are derived by making the assumption that all cells have the shape of cylinders with hemispherical caps using the following equation:
V = &#960; /4 &#215; w2 (L &#8211;w/3)
where L is length and w is width of each cell23,4. Microbial biomass can then be estimated using previously established size-dependent relationships for marine microbial communities24. Generally marine bacteria range in length from 0.1-4 &#181;m, but go up to ~ 8 &#181;m in some locations.
While the filters (0.2 &#181;m) stained with DAPI will only show bacteria, the filters used for the SYBR Gold stain (0.02 &#181;m) contain bacteria and viruses. Measuring the abundance of viruses follows the same protocol as for microbes, however an excitation of 325-375 nm will be used and the emission maximum is at 537 nm (Figure 6B).
In order to generate quantitative data the sampling volume may need to be adjusted depending on the viral abundance in the original sample. The correct volume to filter is best determined empirically for a given body of water. Examples of micrographs containing samples with varying viral concentrations are illustrated in Figure 7.
Results from previous studies suggest that Virus to Microbe ratios (VMR) generally range from 1 to 50 in aquatic systems25-29, and in between 3 and 20, with an average of approximately 6 in coral reef systems (Knowles unpublished data).
Flow Cytometry
In addition to the direct counts and size estimation of the microbial community, assessments of the ratio of autotrophic to heterotrophic microbes via flow cytometry can further be extracted from the collected samples (exemplary flow cytometry output given in Figure 8). To determine the number of total Bacteria cells, samples are stained with SYBR Green I and a photomultiplier tube with a 530/30 bandpass filter is used for detection. A channel for chlorophyll (back to back LP mirrors resulting in a range of 675-735 nm) and phycoerytherin (585/42 bandpass filter) is used to count the abundance of autotrophs in unstained samples. To determine the abundance of heterotrophic microbes, the autotrophic counts from the non-stained portion are then subtracted from the SYBR-stained total count (McDole et al. submitted).
Viral Metagenomics
Viral metagenomics utilizes a culture-independent molecular approach to viral ecology, using total genomic sequence information to determine community structure and function. Metagenomics has been advancing our understanding of the complexity and diversity of viral communities at local17 to global scales30. The recent development of a wide array of bioinformatic tools for analysis of viral metagenomic data has decreased computational bottlenecks, allowing for a more comprehensive view the of the virosphere through the lens of metagenomics.
The methods presented here highlight the isolation and enrichment of viral particles from seawater using a combination of pre-filtration with large pore size nylon mesh to remove debris and cellular material, concentration of samples with TFF (Figure 3) and a subsequent 0.45 &#956;m filtration to remove larger cells. This method produces roughly a 100x&#160;concentrated sample (Figures 5B-5E) which allows us to perform downstream processes in smaller volumes. In order to prevent growth of any remaining microbial cells in the viral lysate and therefore changes in the viral abundance of the sample during the long transit time between field station and laboratory, chloroform is often added to a final concentration of 2% for storage. Following isolation and purification of VLPs, epifluorescence microscopy with nucleic acid dyes such as SYBR Gold are &#160;used to verify the presence and purity of the viral particles (Figures 5B-5E).
Here, we present two viral metagenomes from the Southern Line Island coral reef, specifically, the Starbuck (Star7) and Millennium (CAR9; previously known as Caroline) islands. A total volume of 120 L sample water was collected from each site at a depth of 10 meters and processed as described in section 3.7 and 5.2. The VLPs purified from the cesium chloride gradient centrifugation were 3.3 x&#160;108 particles/ml for CAR9 (Figure 5D) and 2.9 x&#160;109 particles per ml&#160;for Star7 as verified according to the method described in paragraph 4 (Figure 5B). The total amount of DNA isolated from these samples were around 400 ng based on the Nanodrop measurement. DNA was amplified using Phi29 polymerase and sequenced by the Environmental Genomics Core facility in 2011. The characteristics of the sequence data is presented in Table 1. Based on similarity searches using existing databases, the majority (&#62;70%) of the sequences often ended up uncharacterized, i.e., unknown origins and functions. Similarly, the two viromes presented here presented more than 97% of unknown sequences. As a result, non-database dependent analysis was used for analysis and opened up a new arm of research opportunity on the &#8220;dark matter&#8221; in viral metagenomics (Seguritan et al. in press). 
Microbial Metagenomics
The analysis of microbial metagenomes allows for the characterization of the microbial community present and the functional description of these communities. Examples include estimates of the presence of pathogens and virulence5,22 or comparisons between changes in macrobial community structure or available nutrients and abundance of species and their predominant metabolic pathways (Figure 9; Kelly et al.&#160;2014).
Water Chemistry
Water chemistry parameters generally take extensive analytical processing to generate the desired data; samples for DOC analysis will be measured via high temperature catalytic oxidation31,32, particulate organic matter (POM) via isotope ratio mass spectrometry coupled to an elemental analyzer 33-35, and inorganic nutrients by Flow Injection Analysis36,12,37. After a successful completion of all the analyses, information as shown in Table 1 will be available to complement the characterization of the microbial and viral communities. This information can be complimented with stable isotope ratios of organic carbon and nitrogen samples (See Haas&#160;et al.35) and ratios between autotrophs and heterotrophs (McDole et al. submitted).
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/52131/52131fig1highres.jpg" width="600" /> 
Figure 1.&#160;Overview of methods described in the protocol section. <a href="https://www.jove.com/files/ftp_upload/52131/52131fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52131/52131fig2highres.jpg" /> 
Figure 2.&#160;Sample processing rig setup.&#160;The filtration setup introduced here (schematic drawing left, actual picture right panel) is not necessarily required to generate samples but will expedite the process significantly. The setup consists of a backplate, on which the Hatay Niskin units (1) are mounted. On the outlet facing downwards, Hatay Niskin units are connected with silicone tubing to the in-line filter holders (2). These let the sampled water pass through the respective filter into the HDPE sampling vials (3) mounted right underneath. The filtration process is accelerated through pressurized air (4), which and regulated by a pressure gage (5). An overflow basin prevents spilling water during the changing of the HDPE vials or POM filtration. <a href="https://www.jove.com/files/ftp_upload/52131/52131fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52131/52131fig3highres.jpg" /> 
Figure 3.&#160;Tangential flow filter lineup (modified from Thurber&#160;et al.19).&#160;Nylon mesh filtered sample water is pumped from the reservoir (1) via a peristaltic pump (2) to a tangential flow filter (3). The sample water returns to the reservoir (1) along the return line (4) in the absence of backpressure. When backpressure is applied through a hose clamp (5) on the return line, water passes through the filter and is discarded or delivered to the filtrate reservoir (6). Sample water is replaced as it is concentrated out the filtrate line until all the sample water is concentrated in the tubing lines.
<img alt="Figure 4" src="/files/ftp_upload/52131/52131fig4highres.jpg" /> 
Figure 4.&#160;Microscope slide filtration setup.&#160;To evenly distribute DAPI and SYBR Gold stained samples on the respective alumina matrix filter (1), filters need to be placed in between the filter tower (2) and stem of the filtration manifold and fixed with a clamp (4). A pressure regulated vacuum (5) will expedite the filtration process (schematic drawing top, actual picture bottom panel). <a href="https://www.jove.com/files/ftp_upload/52131/52131fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52131/52131fig5highres.jpg" /> 
Figure 5.&#160;Purification and concentration of phage particles.&#160;A total of 1 ml of each cesium chloride density gradient is layered on top of each other prior to leading the pre-treated sample (A). Each layer should be marked outside the tube to facilitate the extraction after ultracentrifugation. Red arrow marks where the tube will be pierced to extract the VLP fraction. (B-E)&#160;Micrographs of viral concentrates: Epifluorescence micrographs of 0.45&#160;&#956;m-filtered&#160;representative viral concentrates of Star7 (B) and CAR9 (D). Large particles including bacterial cells were visible from the 0.45 &#956;m-filtrates from both Star7 (B) and CAR9 (D) samples, whereas only VLPs (white arrows) remain after cesium chloride gradient ultracentrifugation; Star 7 (C) and CAR9 (E). <a href="https://www.jove.com/files/ftp_upload/52131/52131fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/52131/52131fig6highres.jpg" /> 
Figure 6.&#160;Example of DAPI and SYBR Gold micrograph analysis.&#160;Screenshot from the analysis of (A) DAPI and (B) SYBR Gold stained slide example. (A) Length and width (&#181;m) of all highlighted particles (= microbes) can be assessed with the help of an imaging program. Note the aggregation in the center of the image. These clusters will need to be excluded from subsequent analysis. (B) During analysis viruses and bacteria need to be binned by size thresholds into VLP and cellular size ranges (VLP red, cellular green). Note that there are usually some faint VLPs uncategorized by the respective imaging program. These VLPs appearing as faint white dots must be manually counted and added to the automated viral counts. <a href="https://www.jove.com/files/ftp_upload/52131/52131fig6highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/52131/52131fig7highres.jpg" /> 
Figure 7.&#160;Concentrations of SYBR Gold stained samples.&#160;Micrographs of a 0.02 alumina matrix filter containing various numbers of SYBR stained virus samples. Panel A shows a filter containing a suitable amount of seawater filtered, whereas the concentrations on the filter shown in B are too high and in C too low for reliable counts. <a href="https://www.jove.com/files/ftp_upload/52131/52131fig7highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" fo:content-width="6in" src="/files/ftp_upload/52131/52131fig8highres.jpg" width="600" /> 
Figure 8.&#160;Results of flow cytometry analysis output of a reef-water sample.&#160;Panel (A) shows molecular grade water only with yellow-green fluorescent microspheres (0.75 &#956;m), used to verify minimal background with the instrument settings. Panel (B) depicts the output of the reef water sample run with identical settings and layout as in A.&#160;Note that the beads can still be seen in the same location, along with several populations of autotrophs. (C) The same sample as in (B), used to back-gate the flow cytometer, targeting SYBR positive events (autotrophic + heterotrophic count) and minimizing background. (D) The representative reef-water sample stained with SYBR Green I. Using the gating established in (C), total microbe counts were generated, and heterotrophic and autotrophic microbes partitioned by chlorophyll autofluorescence (see McDole&#160;et al.4). <a href="https://www.jove.com/files/ftp_upload/52131/52131fig8highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" fo:content-width="6in" src="/files/ftp_upload/52131/52131fig9highres.jpg" width="600" /> 
Figure 9.&#160;Example of microbial metagenome data.&#160;Patterns between the microbial metagenomic sequence data and site dependent variables. Here, the relative abundance of Synechococcus spp. was positively correlated with the percent cover of coral, whereas Pelagibacter spp. was not (left panels). The metabolic pathway for conjugative transfer was positively correlated with nitrate concentrations, while the metabolic pathway for DNA repair was consistent between all metagenomes (right panels). <a href="https://www.jove.com/files/ftp_upload/52131/52131fig9highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td>Star 7</td>
			<td>CAR9</td>
		</tr>
		<tr>
			<td>Total Number of Reads</td>
			<td>939,311</td>
			<td>591,600</td>
		</tr>
		<tr>
			<td>Number of Preprocessed Reads</td>
			<td>579,664</td>
			<td>360,246</td>
		</tr>
		<tr>
			<td>Mean Sequence Length (bp)</td>
			<td>395 &#177; 131</td>
			<td>451 &#177; 159</td>
		</tr>
		<tr>
			<td>Annotated Sequences (%)</td>
			<td>42,218 (7.3%)</td>
			<td>9,117 (2.5%)</td>
		</tr>
		<tr>
			<td>Unknown (%)</td>
			<td>537,446 (92.7%)</td>
			<td>351,129 (97.5%)</td>
		</tr>
	</tbody>
</table>
Table 1. Data characteristics of two viromes generated from the Southern Line Islands, Starbuck and Millennium. The degree of annotation is based on BLAT using the MG-RAST server.
<img alt="Table 2" fo:content-width="7in" src="/files/ftp_upload/52131/52131table2.jpg" width="700" /> 
Table 2.&#160;Representative results showing data from various sites during one expedition. The here assessed parameters include water chemistry measurements paired with microbial and viral abundances. The table further contains file names referring to the respective metagenomic data for each sampling location. Carbon and inorganic nutrient concentrations are given in &#956;mol/L. <a href="https://www.jove.com/files/ftp_upload/52131/52131table2.jpg" target="_blank">Please click here to view a larger version of this table.</a>

Watch this Scientific Journal Video about Unraveling the Unseen Players in the Ocean - A Field Guide to Water Chemistry and Marine Microbiology at JoVE.com

Unraveling the Unseen Players in the Ocean - A Field Guide to Water Chemistry and Marine Microbiology

Here, we present a comprehensive protocol to assess the organic and inorganic nutrient availability and the abundance and structure of microbial and viral communities in remote marine environments.

Here we introduce a series of thoroughly tested and well standardized research protocols adapted for use in remote marine environments. The sampling ...

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25.0K vues.  San Diego State University. Here, we present a comprehensive protocol to assess the organic and inorganic nutrient availability and the abundance and structure of microbial and viral communities in remote marine environments. 

Vidéo: Unraveling the Unseen Players in the Ocean - A Field Guide to Water Chemistry and Marine Microbiology

A Fish-feeding Laboratory Bioassay to Assess the Antipredatory Activity of Secondary Metabolites from the Tissues of Marine Organisms

Marine chemical ecology is a young discipline, having emerged from the collaboration of natural products chemists and marine ecologists in the 1980s with the goal of examining the ecological functions of secondary metabolites from the tissues of marine organisms. The result has been a progression of protocols that have increasingly refined the ecological relevance of the experimental approach. Here we present the most up-to-date version of a fish-feeding laboratory bioassay that enables investigators to assess the antipredatory activity of secondary metabolites from the tissues of marine organisms. Organic metabolites of all polarities are exhaustively extracted from the tissue of the target organism and reconstituted at natural concentrations in a nutritionally appropriate food matrix. Experimental food pellets are presented to a generalist predator in laboratory feeding assays to assess the antipredatory activity of the extract. The procedure described herein uses the bluehead, Thalassoma bifasciatum, to test the palatability of Caribbean marine invertebrates; however, the design may be readily adapted to other systems. Results obtained using this laboratory assay are an important prelude to field experiments that rely on the feeding responses of a full complement of potential predators. Additionally, this bioassay can be used to direct the isolation of feeding-deterrent metabolites through bioassay-guided fractionation. This feeding bioassay has advanced our understanding of the factors that control the distribution and abundance of marine invertebrates on Caribbean coral reefs and may inform investigations in diverse fields of inquiry, including pharmacology, biotechnology, and evolutionary ecology.

This bioassay employs a model predatory fish to assess the presence of feeding-deterrent metabolites from organic extracts of the tissues of marine organisms at natural concentrations using a nutritionally comparable food matrix.

Marine chemical ecology is a young discipline, having emerged from the collaboration of natural products chemists and marine ecologists in the 1980s with ...

Automated Gel Size Selection to Improve the Quality of Next-generation Sequencing Libraries Prepared from Environmental Water Samples

Next-generation sequencing of environmental samples can be challenging because of the variable DNA quantity and quality in these samples. High quality DNA libraries are needed for optimal results from next-generation sequencing. Environmental samples such as water may have low quality and quantities of DNA as well as contaminants that co-precipitate with DNA. The mechanical and enzymatic processes involved in extraction and library preparation may further damage the DNA. Gel size selection enables purification and recovery of DNA fragments of a defined size for sequencing applications. Nevertheless, this task is one of the most time-consuming steps in the DNA library preparation workflow. The protocol described here enables complete automation of agarose gel loading, electrophoretic analysis, and recovery of targeted DNA fragments. 
In this study, we describe a high-throughput approach to prepare high quality DNA libraries from freshwater samples that can be applied also to other environmental samples. We used an indirect approach to concentrate bacterial cells from environmental freshwater samples; DNA was extracted using a commercially available DNA extraction kit, and DNA libraries were prepared using a commercial transposon-based protocol. DNA fragments of 500 to 800 bp were gel size selected using Ranger Technology, an automated electrophoresis workstation. Sequencing of the size-selected DNA libraries demonstrated significant improvements to read length and quality of the sequencing reads.

This manuscript describes an automated gel size selection approach for purifying DNA fragments for next-generation sequencing. The Ranger Technology provides complete automation of the entire process of agarose gel loading, electrophoretic analysis, and recovery of targeted DNA fragments allowing for high-throughput and high quality next-generation sequencing libraries.

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A Duplex Digital PCR Assay for Simultaneous Quantification of the Enterococcus spp. and the Human Fecal-associated HF183 Marker in Waters

This manuscript describes a duplex digital PCR assay (EntHF183 dPCR) for simultaneous quantification of Enterococcus spp. and the human fecal-associated HF183 marker. The EntHF183 duplex dPCR (referred as EntHF183 dPCR hereon) assay uses the same primer and probe sequences as its published individual quantitative PCR (qPCR) counterparts. Likewise, the same water filtration and DNA extraction procedures as performed prior to qPCR are followed prior to running dPCR. However, the duplex dPCR assay has several advantages over the qPCR assays. Most important, the dPCR assay eliminates the need for running a standard curve and hence, the associated bias and variability, by direct quantification of its targets. In addition, while duplexing (i.e. simultaneous quantification) Enterococcus and HF183 in qPCR often leads to severe underestimation of the less abundant target in a sample, dPCR provides consistent quantification of both targets, whether quantified individually or simultaneously in the same reaction. The dPCR assay is also able to tolerate PCR inhibitor concentrations that are one to two orders of magnitude higher than those tolerated by qPCR. These advantages make the EntHF183 dPCR assay particularly attractive because it simultaneously provides accurate and repeatable information on both general and human-associated fecal contamination in environmental waters without the need to run two separate qPCR assays. Despite its advantages over qPCR, the upper quantification limit of the dPCR assay with currently available instrumentation is approximately four orders of magnitude lower than that achievable by qPCR. Consequently, dilution is needed for measurement of high concentrations of target organisms such as those typically observed following sewage spills.

A Duplex Digital PCR Assay for Simultaneous Quantification of the Enterococcus spp. and the Human Fecal-associated HF183 Marker in Waters

This manuscript describes a duplex digital PCR assay that can be used to simultaneously quantify Enterococcus spp. and the HF183 genetic markers as indicators of general and human-associated fecal contamination in recreational waters.

This manuscript describes a duplex digital PCR assay (EntHF183 dPCR) for simultaneous quantification of Enterococcus spp. and the human fecal-associated ...

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination control. Developments of some techniques aiming to reduce trace metal contamination in the last couple of decades have resulted in concentrations reported now being orders of magnitude lower than those in the past. These low concentrations often necessitate preconcentration of water samples prior to instrumental analysis of samples. Since contamination can appear in all phases of trace metal analyses, including sample collection (and during preparation of sampling containers), storage and handling, pretreatments, and instrumental analysis, specific care needs to be taken in order to reduce contamination levels at all steps. The effort to develop and utilize "clean techniques" in trace metal studies allows scientists to investigate trace metal distributions and chemical and biological behavior in greater details. This advancement also provides the required accuracy and precision of trace metal data allowing for environmental conditions to be related to trace metal concentrations in aquatic environments.
This protocol that is presented here details needed materials for sample preparation, sample collection, sample pretreatment including preconcentration, and instrumental analysis. By reducing contamination throughout all phases mentioned above for trace metal analysis, much lower detection limits and thus accuracy can be achieved. The effectiveness of "clean techniques" is further demonstrated using low field blanks and good recoveries for standard reference material. The data quality that can be obtained thus enables the assessment of trace metal distributions and their relationships to environmental parameters.

Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination ...

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and detritus, and excreting particulate and dissolved compounds, they serve as important agents for benthic-pelagic coupling. Accurately measuring the compounds removed and excreted by suspension feeders (such as sponges, ascidians, polychaetes, bivalves) is crucial for the study of their physiology, metabolism, and feeding ecology, and is fundamental to determine the ecological relevance of the nutrient fluxes mediated by these organisms. However, the assessment of the rate by which suspension feeders process particulate and dissolved compounds in nature is restricted by the limitations of the currently available methodologies. Our goal was to develop a simple, reliable, and non-intrusive method that would allow clean and controlled water sampling from a specific point, such as the excurrent aperture of benthic suspension feeders, in situ. Our method allows simultaneous sampling of inhaled and exhaled water of the studied organism by using minute tubes installed on a custom-built manipulator device and carefully positioned inside the exhalant orifice of the sampled organism. Piercing a septum on the collecting vessel with a syringe needle attached to the distal end of each tube allows the external pressure to slowly force the sampled water into the vessel through the sampling tube. The slow and controlled sampling rate allows integrating the inherent patchiness in the water while ensuring contamination free sampling. We provide recommendations for the most suitable filtering devices, collection vessel, and storing procedures for the analyses of different particulate and dissolved compounds. The VacuSIP system offers a reliable method for the quantification of undisturbed suspension feeder metabolism in natural conditions that is cheap and easy to learn and apply to assess the physiology and functional role of filter feeders in different ecosystems.

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

We introduce the VacuSIP, a simple, non-intrusive, and reliable method for clean and accurate point sampling of water. The system was developed and evaluated for the simultaneous collection of the water inhaled and exhaled by benthic suspension feeders in situ, to cleanly measure removal and excretion of particulate and dissolved compounds.

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Protocol for Microplastics Sampling on the Sea Surface and Sample Analysis

Microplastic pollution in the marine environment is a scientific topic that has received increasing attention over the last decade. The majority of scientific publications address microplastic pollution of the sea surface. The protocol below describes the methodology for sampling, sample preparation, separation and chemical identification of microplastic particles. A manta net fixed on an &#187;A frame&#171; attached to the side of the vessel was used for sampling. Microplastic particles caught in the cod end of the net were separated from samples by visual identification and use of stereomicroscopes. Particles were analyzed for their size using an image analysis program and for their chemical structure using ATR-FTIR and micro FTIR spectroscopy. The described protocol is in line with recommendations for microplastics monitoring published by the Marine Strategy Framework Directive (MSFD) Technical Subgroup on Marine Litter. This written protocol with video guide will support the work of researchers that deal with microplastics monitoring all over the world.

The protocol below describes the methodology for: microplastics sampling on the sea surface, separation of microplastic and chemical identification of particles. This protocol is in line with the recommendations for microplastics monitoring published by the MSFD Technical Subgroup on Marine Litter.

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Controlled-release of Chlorine Dioxide in a Perforated Packaging System to Extend the Storage Life and Improve the Safety of Grape Tomatoes

A controlled-release chlorine dioxide (ClO2) pouch was developed by sealing a slurry form of ClO2 into semipermeable polymer film; the release properties of the pouch were monitored in containers with or without fruit. The pouch was affixed to the inside of a perforated clamshell containing grape tomatoes, and the effect on microbial population, firmness, and weight loss was evaluated during a 14 day storage period at 20 &#176;C. Within 3 days, the ClO2 concentration in the clamshells reached 3.5 ppm and remained constant until day 10. Thereafter, it decreased to 2 ppm by day 14. The ClO2 pouch exhibited strong antimicrobial activity, reducing Escherichia coli populations by 3.08 log CFU/g and Alternaria alternata populations by 2.85 log CFU/g after 14 days of storage. The ClO2 treatment also reduced softening and weight loss and extended the overall shelf life of the tomatoes. Our results suggest that ClO2 treatment is useful for extending the shelf life and improving the microbial safety of tomatoes during storage without impairing their quality.

Here, we describe a protocol for the application of a novel, slow-release ClO2 product that reduces spoilage and extends the shelf life of fresh fruit. The slow-release ClO2 product was added to standard commercial grape tomato packaging and tested against Escherichia coli and Alternaria alternata.

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Extraction of Organochlorine Pesticides from Plastic Pellets and Plastic Type Analysis

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment during manufacturing and transport. Because of their environmental persistence, they are widely distributed in the oceans and on beaches all over the world. They can act as a vector of potentially toxic organic compounds (e.g., polychlorinated biphenyls) and might consequently negatively affect marine organisms. Their possible impacts along the food chain are not yet well understood. In order to assess the hazards associated with the occurrence of plastic pellets in the marine environment, it is necessary to develop methodologies that allow for rapid determination of associated organic contaminant levels. The present protocol describes the different steps required for sampling resin pellets, analyzing adsorbed organochlorine pesticides (OCPs) and identifying the plastic type. The focus is on the extraction of OCPs from plastic pellets by means of a pressurized fluid extractor (PFE) and on the polymer chemical analysis applying Fourier Transform-InfraRed (FT-IR) spectroscopy. The developed methodology focuses on 11 OCPs and related compounds, including dichlorodiphenyltrichloroethane (DDT) and its two main metabolites, lindane and two production isomers, as well as the two biologically active isomers of technical endosulfan. This protocol constitutes a simple and rapid alternative to existing methodology for evaluating the concentration of organic contaminants adsorbed on plastic pieces.

Microplastics act as vector of potentially toxic organic contaminants with unpredictable effects. This protocol describes an alternative methodology for assessing the levels of organochlorine pesticides adsorbed on plastic pellets and identifying the polymer chemical structure. The focus is on pressurized fluid extraction and attenuated total reflectance Fourier transform infrared spectroscopy.

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The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly relevant as a model for the studies of osmoregulatory mechanisms because of its extreme tolerance to low salinity. It is also widely used as a model of settling biology, in particular in relation to antifouling research. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae for studies. A protocol for the all-year-round culturing of B. improvisus has been developed and a detailed description of all steps in the production line is outlined (i.e., the establishment of adult cultures on panels, the collection and rearing of barnacle larvae, and the administration of feed for adults and larvae). The description also provides guidance on troubleshooting and discusses critical parameters (e.g., the removal of contamination, the production of high-quality feed, the manpower needed, and the importance of high-quality seawater). Each batch from the culturing system maximally yields roughly 12,000 nauplii and can deliver four batches in a week, so up to almost 50,000 larvae per week can be produced. The method used to culture B. improvisus is, probably, to a large extent also applicable to other marine invertebrates with free-swimminglarvae. Protocols are presented for the dissection of various tissues from adults as well as the production of high-quality RNA for studies on gene expression. It is also described how cultured adults and reared cyprids can be utilized in a wide array of experimental designs for examining gene expression in relation to external factors. The use of cultured barnacles in gene expression is illustrated with studies of possible osmoregulatory roles of Na+/K+ ATPase and aquaporins.

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

The barnacle Balanus (Amphibalanus) improvisus is a model for studying osmoregulation and antifouling. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae. Here, a protocol for the all-year-round culturing of B. improvisus is described, including the production of larvae. The use of cultured barnacles in gene expression studies is illustrated.

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly ...

Construction of a Low-cost Mobile Incubator for Field and Laboratory Use

Incubators are essential for a range of culture-based microbial methods, such as membrane filtration followed by cultivation for assessing drinking water quality. However, commercially available incubators are often costly, difficult to transport, not flexible in terms of volume, and/or poorly adapted to local field conditions where access to electricity is unreliable. The purpose of this study was to develop an adaptable, low-cost and transportable incubator that can be constructed using readily available components. The electronic core of the incubator was first developed. These components were then tested under a range of ambient temperature conditions (3.5 &#176;C - 39 &#176;C) using three types of incubator shells (polystyrene foam box, hard cooler box, and cardboard box covered with a survival blanket). The electronic core showed comparable performance to a standard laboratory incubator in terms of the time required to reach the set temperature, inner temperature stability and spatial dispersion, power consumption, and microbial growth. The incubator set-ups were also effective at moderate and low ambient temperatures (between 3.5 &#176;C and 27 &#176;C), and at high temperatures (39 &#176;C) when the incubator set temperature was higher. This incubator prototype is low-cost (&lt; 300 USD) and adaptable to a variety of materials and volumes. Its demountable structure makes it easy to transport. It can be used in both established laboratories with grid power or in remote settings powered by solar energy or a car battery. It is particularly useful as an equipment option for field laboratories in areas with limited access to resources for water quality monitoring.

This paper describes a method for building an adaptable, low-cost and transportable incubator for microbial testing of drinking water. Our design is based on widely available materials and can operate under a range of field conditions, while still offering the advantages of higher-end laboratory-based models.

Incubators are essential for a range of culture-based microbial methods, such as membrane filtration followed by cultivation for assessing drinking water ...