Date of Revision: November 7, 2005
Diagram for Water Sample Splits
Global Water Sample
(2 Liters plus volume for algae 10 Liters of
plus inorganic and organic U.Wash.
chemical analysis )
Hawaii Miami (4 L)
200 ml for E.coli culturing
(0.75 L) (2 Liters)
DNA extraction from Water via
(100, 10, 1 ml dilutions on DNA extract Sterivex filtration:
plates) Sterivex filter from LSU (for
detection and typing of HABS (4 samples as per Sterivex filters (1 L x 3).
Bacteroides and assay for attached protocol, 650 ml Filtering is the joint
200 ml for enterococci
Vibrio and Staph virulence minimum) responsibility between LSU and
genes) WHOI. Ownership of the filters
(100, 10, 1 ml dilutions on
Enterococci using IDEXX is to be maintained by WHOI or
500 ml for FRNA coliphages (40 ml) LSU until relinquished. All
and spore tests shipped filters (3 per site) will be shipped
LSU analysis of E. coli and
enterococci using qPCR. This refrigerated to UH 3 flat polycarbonate filters for to WHOI for DNA extraction.
will be based on the DNA subsequent DNA processing, These extracts will then be
extraction from the Sterivex 1.7 ml formaldehyde- Cepheid method for 1 filter pooled and split and shipped to
and QBiogene for second each participating group.
filters (Water volume included preserved water for
in volumes provided to bacterial/virus counts by filter. Extracts to be analyzed
for enterococci and E. coli DNA extracts from Sterivex
WHOI/MIT) epifluorescence microscopy;
analysis. Last filter is for filters to be analyzed for Vibrio
flash frozen and shipped to
archival purposes. Extract and Legionella, plus eukaryotic
Data collect: T, pH, Cond., UH
may be used for and prokaryotic diversity.
Turbidity, NH3 using YSI
probe. Crypto/Giardia analysis as
*250 ml for cultivation of
Total N and P, nitrate, NH3, budget permits. (300 ml)
Vibrios & Staph with colonies
PO4-3, and H2S from water picked, put in glycerol, and
sample. (volume needed?) DNA extract from Sterivex
frozen for shipment to UH
from LSU/WHOI. Analyze
Algal species by microscopy subset for Crypto/Giardia as
*Only when UH
(volume?) (Is this to be fixed? budget permits. Plus
representative present to
- If so – and people are compare with
handle the work)
interested we could ask that 2 Cepheid/QBiogene DNA
samples are taken – one for extraction method.
observation and one for later
FISH?)** RNA extract (250 ml min.)
All to be analyzed on-site for
VOC and SVOCs in separate Norovirus and enterovirus
glass bottles. using Cepheid kits.
Responsible for archiving
** These samples need to be shipped excess.
overnight express to MBL.
Diagram for Soil and Clam Sample Splits
Global Soil Sample
500? g of U.Wash.
XX g frozen for (None)
subsequent mercury DNA extraction from Sediment:
XX g formaldehyde
analysis. 5 ml sediment plus
fixed and stored at -80
C RNAlater (25ml), store at
4C – This has been
DNA Extract split to changed to 1ml sediment +
be provided to Polz 5 ml RNAlater to reduce
from Steward?. cost– we need to notify
Grieg and see if he wants to
split this sample or collect
FISH sediment sample
4 ml sediment plus 12 ml
4% formaldehyde (Note
change in volumes),
store at 4C, ship to WHOI
within 24 hours.
~ 30 gm sediment for
hydrocarbon in glass jars,
Global Clam Sample
(Collect 10 clams per site)
GLOBAL PROTOCOLS FOR WATER SAMPLE COLLECTION
Collection of Global Water Sample
Samples collected in the 10-L carboys are called “global water samples.” Global water
samples to be split into sub-samples by pouring into bottles/containers provided by the
different research groups.
Sterilization of Carboy: Six 10-L rigid polypropylene carboys are available and
additional 10-L collapsible carboys are also available. The collapsible carboys cannot be
autoclaved and so sterilization will occur as follows for all containers.
1. Rinse with 10:1 dilution of nitric acid
2. Rinse with de-ionized water
3. Rinse with ethanol
4. Rinse with local sample water prior to sample collection
The ideal method for sterilizing the rigid polypropylene carboys is as follows:
Autoclave the polypropylene carboys to inactivate microbes. When autoclaving
the spigot must be removed and disinfected separately in bleach (overnight bleaching is
preferred). Rinse spigot with DI water after bleach. The cap of the carboy can be
autoclaved but the cap should not be screwed on the top of the carboy during autoclaving.
Assemble carboy after autoclaving the main vessel and cap and after bleaching the spigot.
Rinse the carboy with 1-2% HCl and allow some of this to run through the spigot and
come in contact with the cap. The purpose of the 1-2% HCl is to destroy any residual
nucleic acid. After rinsing with the HCl, then follow with a distilled water rinse. Upon
sample collection rinse carboy 3 times with some of the local sample water.
If carboys or other filtration materials are to be re-used in the field they should be rinsed
with a small amount of 1 – 2% HCl if possible (about 20 ml for 1L square bottles
including connectors and 100 ml for 10L carboy), and then rinsed well with distilled
water. Prior to filling with sample water, the bottles and carboy should be rinsed with a
small amount of the local sample water.
Note: Please test polypropylene carboys in autoclave (spigot removed) to assure that
they are autoclavable. This test can be done by putting the carboy on a metal tray which
will capture the plastic in the event that the carboy melts. We have conflicting
information concerning whether or not polypropylene can be autoclaved and so to be safe,
let’s test it at LSU.
Global Protocols for WATER Filtration and Subsequent DNA
Extraction of WATER Samples
All groups have indicated an interest in collection of samples for DNA analysis. Through
October 9, filters were collected by LSU on 0.45 um pore size, flat surface filters, and
then frozen dry. This sample collection protocol has since been augmented with the
collection of additional shared filters including Sterivex filters by Hawaii on October 11
(2 per site no lysis buffer added) and by Miami on October 19 (2 per site no lysis buffer
added), October 24 (4 per site no lysis buffer added), October 25 (4 per site no lysis
buffer added) and Nov 2 (2 per site with lysis buffer added). After the conference call
held Oct. 27, 2005 an agreement was reached to add lysis buffer to the Sterivex filters
(and any additional flat surface filters collected by LSU upon collection). The Sterivex
filters that were stored dry were then retrieved and lysis buffer has since been added. The
Sterivex filters collected on October 11 by Hawaii were then brought back to WHOI,
during Jenny’s return. Additional filtration using Sterivex filters is to follow. Prior to the
return of the Miami group on November 13 the balance of the Sterivex filters will be
shipped to WHOI. WHOI will then extract the DNA from these filters and then ship the
DNA extracts back to each corresponding laboratory.
Protocol For Flat Filters Processed by LSU
Sample Collection on Flat Surface Filters: For consistency let’s continue using the
same filters that have been used by Aixin Hou since the beginning of sample
collection. These filters are 0.45 um in pore size and are consistent with standard
sample collection protocols for bacteria using membrane filtration. These filters were
originally frozen dried and stored.
DNA extraction of Flat Surface Filters: Candidates include: Hot detergent/bead
beating/CTAB/phenol+chloroform (Gast). Any other recommendations? Some had
recommended methods for sediment/soils (e.g. MO BIO soil kit and Gentrasystems
kits). The idea is to provide splits of the extracts to each of the laboratories assuming
that enough is available.
Overview: The sterivex filters are 0.22 micron capsules that are routinely use by WHOI
for their Mt Hope Bay bacteria and eukaryote (3-domain – including Archaea) work. The
set up that allows the filtration of water from 4 one liter bottles at a time (using a vacuum
pump) onto several Sterivex filters. The Sterivex filters do clog eventually (usually
between 200 & 400ml but up to 3L for very clean water) and that's when filtration is
stopped and the sample volume is recorded. See picture below. There are two varieties
of Sterivex filters available, PVDF and PES. PVDF has lower binding and so these are
preferred for virus work where the filtrate will be concentrated further.
WHOI/MBL/MIT (Gast/Zettler) have provided 15 1L bottles and the filtration
apparatus. These, along with the large barrel syringes will need to be washed, acid
treated, and autoclaved prior to each sampling. If these are to be reused in the field, the
1L bottles, connectors and barrel syringes will need to be as a minimum rinsed with 1 -
2 % HCl. WHOI has included bags for sterilization of the syringes. Each group to
provide their own Sterivex filter. Ordering information is as follows: Millipore Sterivex-
GP Filter Unit with Male Luer-Lok cat no. SVGPL10RC - which are the PVDF variety.
Each group will also need Cole-Parmer Cat no. A-30526-08 Male female luer cap
polycarbonate to seal the ends.
Protocol for Filtering Water Samples through 0.22 micron Sterivex Filters: Run
filtration. The number of Sterivex filters processed per site has varied from 2 to 4. After
the October 27, 2005 an agreement was reached to process 3 Sterivex filters per site. The
extracts from these three filters will be pooled and then split into 4. (One split for LSU,
one for Hawaii, one for Miami, and one for WHOI)
Assembly of Sterivex Filtration System*:
1. Clean area for Sterivex Filtration System, (SFS) setup. Attach PVC tubing to
input connector of a 10L carboy and attach the outport to a vacuum pump. We
like to put an in-line filter between the outport and the pump to prevent pump
2. Wearing a clean set of gloves, carefully attach the sterile female/female luer-lock
adaptors to the pipet-end in the stopper. Everything should be sterile from this
3. Label each sterivex filter accordingly. Then attach the male end of the filter to the
top of the female adapter.
4. Attach the 140 cc syringe barrel to the female end of the sterivex (see photo on
5. Fill the bottles to the 1 liter mark. While keeping plenty of paper towels at hand,
quickly invert the 1L bottle of water onto the syringe barrel and secure the bottles
to the board .
6. After all four manifolds are set up, turn on vacuum and open all red t-valves.
7. Let the filtration run until completion or the filter clogs. In the event that the later
occurs, close the t-valve and remove the syringe barrel from the sterivex. Pour
any remaining water into a graduated cylinder and record actual amount filtered.
It is essential that the real amount of water filtered be recorded. This is especially
important for samples subsequently analyzed via qPCR.
Collection of Sterivex Filter**:
1. Reopen the t-value to continue filtration. Run the manifold until the sterivex filter
is dry, then shut off red t-valve.
2. Remove sterivex filter and cap the bottom, or male end, with a blue end cap.
Sometimes it is necessary to flick the sterivex with your wrist to get the final
drops out before capping.
3. Add 2mL, or one tube, of Puregene Lysis Buffer to the Sterivex filter using a
sterile transfer pipet.
4. Cap the top or female end of the Sterivex with a blue end cap.
5. Seal the connection at each end of the Sterivex filter with parafilm and place into
a sterile Whirlpak bag.
6. These filters should be stored on ice for shipping purposes and frozen at –20 C if
not shipped the same day. (Currently stored at -80 C at LSU)
* A second set of hands can aid in cleaning up spills from inverting the 1L bottle.
** Ideally, one person should handle the sterivex filter while a second person helps by
opening blister packs for the caps and Lysis Buffer. This can be done by one individual
with a bit more difficulty.
Figure A: Sterivex Filtration System with Details for Sterivex Filter Connections.
DNA Extraction from Sterivex Filters using the Puregene Kit
(Method provided by Linda Amarel Zettler)
Gloves, parafilm, Autoclaved 2 mL microfuge tubes
Puregene kit (Puregene cat no. D-7000A) from Gentra
Lytic enzyme (Puregene cat no. D-60K4)
Note: Lytic Enzyme is a lyticase similar to Zymolase that breaks the 1-3 beta
glucose linkages in bacterial cell walls. It is suspended in a proprietary glycerol
solution at a concentration of 4000 U/mL. Alternatively, Lysozyme (50mg/mL)
made up in autoclaved water can be added at 67ul per sample.
Proteinase K (20 mg/ml) from the kit
Velcro strips or wire strips to attach filters to the carousel in the hybridization oven
Hybridization oven with rotating carousel (37-650C) or equivalent incubator with
1. Take samples out of dewer and let them thaw (~20min).
2. Add 10 ul Lytic Enzyme.
3. Incubate @ 37C for 30min on rotating carousel (change to 65C when filters are
4. Add 10 ul Proteinase K from refrigerator.
5. Vortex @ max speed for 10 seconds.
6. Incubate @ 65C for 1hr.
7. Attach a 3 cc syringe to the top (female end) of sterivex filter and use the plunger
to push the liquid into a sterile 2 ml labeled microfuge tube. If necessary attach
and repeat to push full volume out.
8. Subdivide this volume into 3 tubes (A,B,C) @~700ul per each
9. Cool to RT ~20 min
10. Add 233ul (per 700ul) of Protein Precipitation Solution (from kit) to each tube.
11. Vortex 20 sec to mix ppt solution with cell lysate.
12. Incubate on ice 15min.
13. Centrifuge at max speed 5min.
14. Pipette supernatant (from protein pellet) into tube with 1 volume 100%IPA.
15. Invert 50 times to mix.
16. Centrifuge 5min @max speed.
17. Pour off supernatant and drain on a paper towel. Add 700 ul of 70% ethanol and
invert the tube several times to wash the DNA pellet.
18. Centrifuge at maximum speed for 5 minutes. Pour off supernatant. Invert and
drain the tube on a paper towel. Air dry for ~20 minutes until dry.
19. Resuspend each pellet in 10 ul of hydration buffer provided with the kit. This will
give a total of 30 ul of concentrated DNA per filter sample.
Given experiments conducted on Oct. 9 on samples C5 and C7 by Jenny Rocca of WHOI
on-site at LSU, the following can be anticipated with respect to DNA.
The Puregene kit yielded 4 - 6 ug/L. Introducing a "pre-spin" prior to concentrating the lysate with PEG
precipitation reduced the yield but may increase purity. Spectra obtained with the Puregene kit displayed a
distinct trough at 230 nm, which is indicative of relatively pure DNA. The UV method appeared to greatly
overestimate the DNA concentration, which is a common bias in UV quantification of nucleic acids.
Despite the nice UV spectra the Puregene generated samples were relatively brown. Subsequent work will
focus on substituting PEG for isopropanol in the Puregene protocol in an effort to remove the brown hue.
Additional work by WHOI group has shown that 1:100 dilution best for DNA after ethanol ppt. At this
dilution the general primer was able to pick up all 3 domains.
GLOBAL PROTOCOLS FOR SOIL SAMPLE COLLECTION AND
Collection of Global Sediment Sample
A container large enough to hold the sediment needed by all groups is needed. This
container should have a wide mouth to facilitate mixing and transfer of the sediment to
sub-sample container jars for each group. This container should be cleaned in a fashion
similar to the carboy used to hold the water sample.
WHOI/MBL/MIT and Hawaii have indicated an interest in sediment samples for
subsequent microbe analyses. Below is the proposed protocol for samples to be
preserved in formaldehyde.
Sediment samples were collected on XXX by the LSU. These samples were processed
by Aixin Hou as follows:
The plastic columns for sediment cores were pre-washed by acid, and the
containers for the surface sediments were individually wrapped sterile cups.
Those samples are now kept at -32oC. If these sediment samples are good for
measuring hydrocarbons and fecal sterols, Aixin can thentransfer them to the HC
Chris Sinigalliano mentioned that Harry Roberts of LSU also collects sediment samples
and that sample collection trips for sediments can be possibly “piggy-backed”.
Summary from Chris Reddy concerning WHOI Needs:
Long and short - from Chris's standpoint, 2 grams or so of the sediments collected for Hg
(or other metals?) would be suitable for his purposes in measuring hydrocarbons and
fecal sterols. The care taken regarding contamination of the "Hg samples" should be
sufficient for the hydrocarbons as well.
To make use of these samples, it would need some transfering to his containers with his
(Question: Chris -or Chris - or Aixin; would it be desirable for Chris Reddy to come
down to do the transferring of current sediment samples to the HC clean containers?)
He also would be able to come down on the 11th or 12th for collecting more samples.
(Question: If the current "Hg samples" are suitable for the first set of HC analyses, is a
second sampling desired? IF so, would a second sampling in November be preferred, or
would a second sampling some months down the road be preferred?)
I think that the people who are going to be using sediments need to consider such
[the following is from Chris Reddy to me]
I am still interested in analyzing surface sediments from Lake Pontchartain for petroleum
hydrocarbons and fecal sterols following Hurricane Katrina. Our objectives would be:
1. To analyze samples and provide data at the highest quality, which would be available
to the whole scientific community.
2. To determine current levels of petroleum hydrocarbon as a means to gauge the extent
and release of oil during and after the hurricane. This data would also be compared to
previous work at Lake Pontchartain and other urban areas.
3. To determine current levels of fecal sterols and potentially other sewage-derived
organic compounds, such as linear alkyl benzenes. This data would be particularly useful
to efforts related to the abundance and distribution of bacteria and viruses in the water
I believe from yesterday_s phone meeting, that surface sediments from the top 1 cm were
collected on October 9 or 11. These samples (~50 g) were collected for mercury and
other trace metals and are currently frozen in storage. I am assuming that they were
collected with standard _trace-metal-clean_ techniques. Of course, the latter is different
than collecting and handling samples for organic compounds.
Based on this available knowledge, I envision at least two scenarios regarding sediment
samples for organic analysis.
I. Subsample the _trace metal_ samples
a. The samples have been collected and hence they provide the earliest record of the
sediment characteristics following the hurricane. With time after the event, the sediment
may be redistributed and some of the organic compounds, especially fresh petroleum,
will begin to weather. Hence, these samples provide the best snapshot_.
b. There appears to be more than enough material from the top 1-cm (50g).
Organic analysis only needs 1 to 3 grams of material.
a. They were not sampled with organic-clean techniques. Simple items like bug spray and
the exhaust from the sampling boat can be huge factors in sample cleanliness. However,
it is my experience that efforts to collect material for trace metals often yield reasonable
results for organics. It will be important to determine what material are the sample
containers. Some plastics contain very high organic backgrounds.
b. These samples still would need to be sub-sampled and shipped back to Woods Hole.
II. WHOI scientists fly down on or about Nov 11/12 and collect samples
1. Organic samples can be collected cleanly and earlier than planned.
2. Samples could also be collected and preserved for Linda Amoral Zettler_s group.
1. It would be an expensive and time-consuming process. If we were to go, we would
bring three people. However, we would only need a boat handler and perhaps a mate. We
could easily handle the sampling and preservation, etc...
2. Logistics need to be planned very quickly. Important issues like boat availibility, type
of boat, sediment collection gear, and travel and housing have to be considered soon.
Protocol for Preservation and Preparation of Sediment Samples
(Provided by Linda Amaral Zettler)
4% Formaldehyde (Diluted from 37% stock using filtered and autoclaved seawater or
fresh water depending on the location of the sediment collection)
Phosphate-buffered Saline Solution (PBS) -
-Dissolve the following into 800mL water:
-adjust pH to 7.4 with HCl and bring volume to 1L
-autoclave for 20min on the liquid cycle
-store at room temperature
Centrifuge – we used a Sorvall RC-3B at 4oC & 9200 rcf with 50mL tubes adaptors
25 mL sterile pipettes and pipette bulb
Sterile collection spoons – Sterileware 2.5mL (Sterileware: Fisher no. 14-429B)
50mL centrifuge tubes
Disposal containers for dry and liquid formaldehyde waste
1. In the lab, dispense 12mL aliquots of the 4% formaldehyde into 50mL centrifuge
tubes. Create one aliquot for each sediment sample and pre-label each tube for
more efficient field collecting.
2. In the field, collect 4 mL of sediment from the VanVeen grab (or other sediment
sampler) using the sterile spoon. Immediately resuspend the sediment in the 12mL
of 4% formaldehyde and put on blue ice or refrigerate.
3. In the lab, centrifuge the samples at 9200 rcf for 10min and pipette off the
formaldehyde into a properly labeled waste container.
4. Add 12mL PBS to the sediment and resuspend using a sterile spoon.
5. Centrifuge the samples at 9200 rcf for 10min and pipette off waste into waste
6. Repeat steps 4 and 5.
7. Add 12mL of 1:1 PBS/Ethanol to the sample and resuspend using a new sterile
8. Parafilm the lids of the centrifuge tubes and store at –20oC until further use.
GLOBAL PROTOCOLS FOR CLAM SAMPLE COLLECTION AND
Collect 10 clams from each of the 9 sites around the lake. Clams are to be placed in
sterile Whirlpak bags upon collection.
1. Scrub external surface of the clam with a brush. If possible, the brush should first
be immersed in 70% isopropyl alcohol.
2. Rinse the clams under running tap water.
3. With a sterile clam shucker open the clamr aseptically
4. To open the clam place the clam shucker into the hinge portion of the oyster and
pry it open.
5. When open, collect the clam liquor by using a 5 ml serological pipet
6. Pool the liquor of the 10 clams but separate the liquor by sampling site
7. The shucked clam meat (with no liquor) should be placed into a sterile whirl-pak
8. Please put one clam in each bag and DO NOT POOL THE CLAM MEAT.
9. Immerse the clam shucker in 70% isopropyl alcohol then flame between each
clam sample to prevent cross contamination.
PROTOCOLS SPECIFIC TO EACH RESEARCH GROUP
Protocols for LSU Samples
1. E. coli and enterococci via culture using standard EPA methods. Dilutions used
include 100 ml, 10 ml, and 1 ml. 10 colonies transferred from each plate onto
2. E. coli and enterococci analysis using molecular methods. Is there a standard
3. Filtration of samples through Sterivex system (500 ml x 3). Filtering is the joint
responsibility between LSU and WHOI. Ownership of the filters is to be maintained
by LSU until relinquished to WHOI. WHOI will be responsible for extracting the
DNA, pooling the extracts, and splitting for subsequent shipment to the various
4. T, pH, Cond., Turbidity, NH3 using YSI probe.
5. Total N and P, nitrate, NH3, PO4-3, and H2S from water sample.
6. Algal species by microscopy.
7. VOC and SVOCs in separate glass bottles.
Protocols for Miami Samples
Samples collected for HABs
“ABCD” sample code included. See file called “Sample_Labels…..doc” for an explanation of the ABCD
1. Put around 100 ml of sample water in a purple capped sample cup. Leave
at room temperature at low light. Do not freeze.
ABCD = HNXX (H indicating a sample for HABS and N indicating no pre-processing)
2. Put 8-10 drops of Lugols solution from a dropper bottle into a purple
capped centrifuge tube and then add 50 ml of sample water. Keep in
darkness. Do not freeze.
ABCD = HLXX (H indicating sample for HABS and L indicating that addition of Lugols solution
was part of the pre-processing)
3/4. Filter around 200 ml of sample water onto a 25 mm GF/F filter.
Do this on 2 replicate filters. Fold the filter over twice with the sample
on the inside and place in the 8 ml vials and freeze.
ABCD = HGXX-a or HGXX-b (H indicating sample for HABS, G indicating that sample was
filtered through GF/F, and “a” corresponds to the first filter and “b” corresponds to the second
Samples collected for Microbes
1. Three polycarbonate filters per site. One filter in contact with lysis solution as per the Cepheid
protocol. This extract will be analyzed for Enterococci and E. coli using Cepheid reagents.
Another filter to be stored at -80C for archiving. Third filter to be extracted for DNA using
Qbiogene fastprep kit if Cepheid lysis method found to be inadequate. Extracts from these filters
may be used for Crypto/Giardia analysis, budget permitting.
2. Analysis of water sample for enterococci via IDEXX chromogenic substrate method, in duplicate.
Sample volumes used will be 10 ml diluted up to 100 ml. ABCD = IXXX (I=IDEXX)
3. RNA extraction using virus sorption followed by analysis for Norovirus and enterovirus once
Chris/Maribeth arrive in mid-October. Two extracts to be performed per site with the extracts
pooled into one tube. The filters used in the virus sorption method will be archived, in case they
can be used later.
ABCD = KRXX (K = Katayama based method and R for RNA)
4. Assist with filtration of Sterivex filters
Concentration Step for Viruses (modified after Katayama et al. 2002). Please check procedure against
original paper, copy provided to Shiba and Chris/Maribeth.
a. Setup medium size filtration unit with 1 L sidearm flask
b. Use negative charge membrane filter: 47 mm diameter, 0.45 um
c. Filter 100 to 200 ml of sample. Record volume added.
d. Rinse with 50 mL of 0.5 mM H2SO4, pH 3
i. Dilute 50mM H2SO4 to make (1:100)
e. Measure the sample volume
a. Add 12.5 uL of 50mM H2SO4 into 50 mL sidearm flask
b. Add 12.5 uL of 100x TE buffer in to the sidearm flask
c. Replace the funnel from 1 L sidearm flask to 50 mL sidearm flask
d. Add 2.5 mL of 1mM NaOH into the funnel
i. Need to make
e. Measure the sample volume
f. Keep the filter in a Petridish and put in freezer (ABCD=KDXX)
a. Add the filtrate sample into the Amicon concentrator spin filter. Amicons can be ordered
from Millipore Amicon ultra centrifuge filter devices, UltracelTM low binding
regenerated cellulose for volumes less than 15 ml, 10,000 MWCO. Cat UFC901008.
b. These spin filters reduce the volumes from 15 ml to 200 to 250 ul.
c. Centrifuge for 10 minutes at 10,000 x g
d. Discard the water that does not contain the viruses (read manufacturer instructions.
e. Measure the concentrated sample in bottom of the tube using a pipette. Should be
roughly 200 to 250 ul.
D. RNA Extraction (Using QIAamp Viral RNA Mini Spin Protocol) Below is a recap of the
I. Incubation (RUN IN DUPLICATE, AT END MIX THE RNA EXTRACT IN ONE
CENTRIFUGE TUBE AND SPLIT IN TWO. ONE SPLIT USED FOR ANALYSIS AND
ANOTHER ARCHIVED. Also please check instructions below against the instructions
provided in the kit. Copy provided to Shiba and Chris/Maribeth.)
a. Add 140 uL of the concentrated sample into a 1.5 mL micro centrifuge tube
b. Add 560 uL of Buffer AVL into the tube
i. need to prepare for the first time
ii. Heat it up (60 ~ 80ºC) and cool it down to room temp if it there is precipitation
c. Vortex the tube for 30 seconds
d. Leave the tube for 10 minutes at the room temperature
e. Add 560 uL of ethanol (96-100%) in to the tube
f. Leave the tube for 10 minutes at the room temperature
a. Add 630 uL of the sample into the QIAamp spin column (in a 2 mL collection tube) from
the 1.5 mL micro centrifuge tube
b. Centrifuge the spin tube at 6000 x g (or 8000 rpm) for 1 minutes
c. Place the spin column in to a new clean 2 mL collection tube.
d. Discard the tube containing the filtrate
e. Repeat (a ~ d) until the all sample in the micro centrifuge tube is transferred
i. RNA is on the filter now (the filter is your sample)
a. Place the spin column in a new clean 2 mL collection tube
b. Add 500 uL of Buffer AW1 in to the spin column
c. Vortex the tube for 1 minute
d. Discard the filtrate
e. Add 500 uL of Buffer AW2
f. Vortex the tube for 3 minutes
g. Discard the filtrate
h. Place the spin column in a new clean 2 mL collection tube
i. Vortex the tube for 1 minute
j. Discard the old collection tube containing the filtrate
a. Place the spin column in a new 1.5 mL micro centrifuge tube
b. Add 60 uL of Buffer AVE in to the spin column
c. Leave it for 1 minutes for incubation
d. Centrifuge at 6000 x g (8000 rpm) for 1 minute
e. Discard the spin column (including filter)
f. The filtrate in the micro centrifuge tube is your sample. ABCD = KRQX-a or KRQX-b
(where K = Katayama method, R = RNA extracted, Q = Qiagen kit used. Label “a” for
sample given to Chris/Maribeth for analysis and “b” for archive.) Remember that
duplicates will be run and it would be best to recombine the two extracts and then split
again to assure homogeneity.
g. Store the tube at -20 to - 70ºC
Protocols for Hawaii Samples (Water component only)
1. Total counts of bacteria and viruses: Add 1.7 ml of each sample in duplicate to
cryovials containing 0.1 ml formaldehyde (cryovials with formaldehyde will be
shipped from UH). Store samples at -80°C, ship on dry ice to UH as soon as
2. Analyses for FRNA coliphages, spores, and Bacteroides: Fill sterile 500 ml bottles
with water from each station. Keep somewhat cool if possible, but not cold (>10° C,
PLEASE DO NOT ICE OR REFRIGERATE). Ship to UH as soon as possible (next
morning after sampling).
3. Bivalve processing: Collect 10 clams (or oysters) from each site.
1. Collect 10 clams (or oysters) from each site
2. Scrub external surface of the oyster with a brush. If possible, the brush should
first be immersed in 70% isopropyl alcohol.
3. Rinse the clams under running tap water.
4. Open the clam aseptically with sterile shucker
5. When open, collect the any liquor by using a 5 ml serological pipet
6. Pool the liquor of the 10 clams in WhirlPak bag, but separate the liquor by
7. The shucked clam meat (with no liquor) should be placed into a sterile whirl-pak
8. Please put one clam in each bag and DO NOT POOL THE CLAM MEAT.
9. Immerse the shucker in 70% isopropyl alcohol then flame between each
sample to prevent cross contamination.
1. Collect as many clams as practical at each site (up to 10).
2. Store clams on ice during collection trip
3. Freeze upon return to the laboratory (-80°C preferred, -20°C O.K.)
4. Ship whole frozen clams on dry ice to UH
Protocols for WHOI/MBL/MIT Water and Soil Samples
1. Sterivex filter (frozen in Puregene lysis buffer, shipped to WHOI). Three filters
per site with the exception of the samples collected by Grieg on October 11 which
corresponded to 2 filters per site and the samples filtered by Chris/Maribeth/Shiba
on October XX and XX which corresponded to 4 filters per site. Protocol for
sterivex is in above section. Sterivex filters will be frozen in extraction buffer by
local Louisiana group.
ABCD = SDPX (S=Sterivex, D=DNA extraction, P=Puregene method)
2. Sediments for DNA extraction – 1 ml surface sediment into 50ml centrifuge tube
plus 5ml of RNAlater. Store at 4C or –20C. We will use the Power Soil kit by
MoBIO to process these samples.
3. Sediments for FISH – 4 ml surface sediment into 50 ml centrifuge tube plus 12 ml
4% formaldehyde. See attached doc (Formaldehyde_LAZ_edit.doc) for formalin
sample processing (this will be done at MBL or by Jenny for Oct samples if
equipment is available).
4. Sediments for hydrocarbon analysis – 30 gm surface sediment into glass jars.
Collect sediment as soon as it arrives on deck, and try to keep clear of exhaust and
gasoline fumes. Cap jar immediately and keep cool or frozen.
These are for C. Reddy. We should be receiving 48 jars and 27 pre-cleaned
sampling spoons.The jars are glass with Teflon-lined lids and have a 60-ml
capacity. We would appreciate that the jars are filled halfway to the top and stored
ideally in a freezer but kept cool is ok. They can be shipped with a few ice packs
and do not need dry ice. No preservation is needed. Do not worry about being
sterile but wearing gloves is encouraged. All you need to do is scoop some of the
surface mud (top 1 cm or so) with the spoons, add to the jars, and cap quickly. We
would like two samples from each sample location so we have provided you with
enough for 24 locations. You can use the same spoon for sampling for each site.
Each spoon has been pre-cleaned and all you need to do is remove the aluminum
foil. Do not recycle the spoons. You do not need to send back the spoons but can
if you wish. Please note in the chain-of-custody any samples that smell of oil.
Feel free to label the samples in any code or system that you wish. You can use
labels, which are enclosed, or write on the tops of the lids with a Sharpie.
Protocols for U.Washington Water and Soil Samples
1. To be determined.