Laboratory

The HauriLAB is located in the International Arctic Research Center at the University of Alaska Fairbanks. Our mission is to produce high-quality inorganic carbon data from open ocean and coastal environments.  Our lab is outfitted with an Apollo DIC Analyzer AS-C6, and Kongsberg HydroFIA pH, and HydroFIA TA instrumentation for Dissolved Inorganic Carbon (DIC), pH, and Total Alkalinity (TA) discrete seawater analysis.

High-quality data requires Best Practices during sampling, storage, and analysis.  For example, a sample can quickly become compromised by gas exchange when it is inappropriately handled. To avoid contamination and achieve high accuracy and precision, we follow strict sampling and analysis protocols [Dickson et al., 2007] and have adjusted the Best Practices for inorganic carbon sampling in coastal regions as outlined below.

These Best Practices are the baseline for our lab policies and need to be strictly followed by all our collaborators.  As such, if samples are not collected following the best practices outlined below, we may be unable to perform analysis in order to avoid damage to instrumentation and to ensure that continued high-quality data is generated in our lab. Sampling methods that have not been fully vetted in the peer reviewed literature and adopted as standard Best Practices need to be adequately tested in collaboration with our lab.

We welcome new collaborations, so please contact us if you have new project ideas, upcoming projects that involve inorganic carbon sampling/analysis, or if you are interested in our ongoing projects.  

Email us to schedule an appointment and discuss further details on sampling, instrumentation usage, costs etc..

Cheers,

Claudine Hauri, chauri@alaska.edu, (907) 474-7059

Brita Irving, bkirving@alaska.edu, (907) 474-5966

BEST PRACTICES

Summary of most important points

  • Seawater must be free of sediment: if sampled in coastal areas, follow filtration technique by Bockmon and Dickson [2014] with a filter pore size of 0.45μm.
  • The following water volumes are required for
    • DIC analysis: 10 mL (5mL sufficient for porewater samples)
    • TA analysis:   250 mL
    • pH analysis:   175 mL
  • Samples for DIC and pH must be analyzed as soon as sample bottle is opened
  • The analysis will require a Certified Reference Material (CRM) at the beginning and end of the day, in addition to at least one for every eight samples analyzed. CRM’s can be ordered from Andrew Dickson’s lab (co2crms@ucsd.edu).
  • For pH and TA analyses sample salinity is required. If salinity is unknown, an estimated salinity value will be used during analysis and correct pH and TA values can be calculated using the recorded data together with the known sample salinity.
  • Note: To calculate complete inorganic system parameters (such as with CO2SYS) you will need sample water temperature, pressure or depth that the sample was collect from, salinity, and nutrients (an error of 1-5% can be introduced if nutrient data is unknown).

Best Practices for Open Ocean Sampling

  1. Planning of inorganic carbon sampling

It is best practice to analyze all three variables from the same bottle, which will require 500 mL. DIC and pH will be analyzed at the same time, TA within a few days.

  • Bottles

Sample bottles must be 500 mL Borosilicate 3.3 Glass bottles that meet ASTM E-438, TYPE-1, CLASS- A Standard – ideally with a stopper. Bottles can be purchased through Fisher or VWR. The following water volumes are required for:

  • DIC analysis: 10 mL (5 mL sufficient for porewater samples)
  • TA analysis:   250 mL
  • pH analysis:   175 mL

  • Water sampler size

In order to collect your samples properly, you will need a water sampler that can collect several times the volume of water needed to completely fill the sample bottles. This is because the collection method involves overflowing the sample bottle with two full exchanges of water to collect a clean sample. The sample must not include the last water to come out of the sampler, which will have been exposed to air as the sampler is emptied into the sample bottle and will thus be contaminated. To collect a 500 mL sample, the smallest acceptable water sampler would be a 2 L Niskin if you are exceptionally careful in collecting the sample, but a 2.5 L water sampler or larger would be preferable to ensure sample integrity.

  • Nutrients

Depending on project objectives and in order to make the highest quality calculations, you will need concentration data for phosphate ([PO43–]) and silicate ([SiO44–]). Not having nutrient data can introduce an error on the order of 1–5% or possibly higher, depending on nutrient concentration in the calculated pCO2 values.  Please contact Mette Kaufmann (mrkaufman@alaska.edu) to have your nutrient samples analyzed at IARC.

2. Preparation prior to drawing your samples

  • Prepare a saturated solution of mercuric chloride (HgCl2) in advance

Read the SDS for mercuric chloride. The saturation of mercuric chloride is 7.4 grams per 100 mL of water, this is the minimum ratio you can use. NOAA PMEL suggests a 1:10 ratio for a saturated solution; e.g. 10 grams mercuric chloride per 100 mL water should be sufficient for 200 samples. Ensure that your pipette (or preferably repipettor) is capable of dispensing 200 µL properly. Make sure that the solution stays saturated at all times by checking to see that crystals remain in the bottom of the bottle.

Inorganic carbon samples must be preserved using mercuric chloride, which is a toxic chemical and thus requires special handling. For information on shipping samples with a very low concentration of HgCl2 in them, please see the links on PMEL’s website for information about shipping and small quantity exceptions.

Information about mercuric chloride for sample preservation, pipette or repipettor for dispensing mercuric chloride solution, and water samplers can be found on PMEL’s website.

  • Label the sample bottles in at least two locations

Bottle numbers can rub off, which results in loss of data for any samples that cannot be matched to your sample log. Each bottle should be labeled with a unique number (e.g. A1 to A20, PS-1 to PS-20, etc.) and be placed back in the storage crates sequentially. Use DecoColor paint markers for labeling, since Sharpies (and similar) do not reliably remain clear and are easily rubbed off.

  • Grease the stoppers

Apply a thin strip of Apiezon L grease around the bottom of each bottle stopper (see picture at right for proper greasing method). After sampling, you will insert a stopper into the neck of the sample bottle and twist to spread the grease evenly and form a good seal. Please be aware that this grease can contaminate samples for dissolved organic carbon analyses, so care should be taken to prevent this grease from getting onto shared water sampling equipment.

  • Soak your noodles

Soaking sampling tubing (“noodles”) in a bucket of clean water before the first sample and between subsequent samples helps to prevent bubbles from forming in the noodle during sampling. Remember: bubbles are the enemy of high-quality dissolved gas samples! Noodles may be either a single piece of tubing or consist of a narrower piece of flexible tubing inserted into a larger diameter piece of rigid tubing, and held in place with a cable tie. For single-piece noodles, we recommend marking the noodle so that the same end is placed on the water sampler each time. For two-piece noodles, the rigid tubing goes into the sample bottle and the smaller, more flexible end slips onto the water sampler hose-barb or nipple.

  • Prepare your log sheets

Prepare log sheets with all necessary information including dates and times, locations and depths to be sampled, geographic coordinates, bottle numbers, comments, etc (see above).

3. Sample drawing procedure

  • Draw samples immediately after bottles come aboard

Dissolved gases must be sampled before other less sensitive samples such as nutrients and salinity are collected. The correct order of sampling is oxygen first, then dissolved inorganic carbon parameters (pCO2, pH, dissolved inorganic carbon, and alkalinity).

  • Check the water sampler for leaks

Before opening the air valve on a bottle, open the sample valve by pushing the outer petcock ring in. If water begins to flow out or if water is dripping steadily around the end caps, the end caps did not seat correctly when the sampler closed, and the sample is compromised. Check that the tubing between the end caps is in good condition and attached correctly. If no leaks are observed, close the petcock/stopcock after checking for leaks.

  • Fill sample bottle

Attach the designated end of the Tygon tubing to the stopcock/petcock of the water sampler. Insert Tygon tubing to the bottom of the sample bottle, open air valve, open water valve, and begin water flow. Invert the bottle over the tube to rinse the bottle carefully with the sample water, moving the tubing to eliminate any air bubbles on the bottle walls. It is critical to prevent exposure of the sample to air bubbles. Slowly right the bottle and begin to fill, pinching the tubing (if necessary) to control the influx of bubbles. Allow the bottle to fill completely and to overflow at least one full volume. Bend and pinch the tubing to stop the water flow while the tubing is still touching the bottom of the sample bottle. Then withdraw while the tubing is still bent and pinched—this creates a ‘calibrated’ headspace of ~1% to allow for sample expansion. Check the bottle for any bubbles; if you see any, discard the sample and redraw.

  • Carefully add 200 µL of HgCl2 with pipette or repipettor to 500 mL sample bottles

do not submerge pipette tip in sample.

  • Insert greased stopper into neck of bottle and twist to form a good seal. There should not be any streaks visible in this greased seal. For cold water sampling, keeping the grease in a thermos or cup of hot water may make grease application easier.
  • Seal bottle with rubber band and collar

This is critical for proper storage. Follow provided instructions and pictures below.

  • Place the whole collar through the middle of the rubber band (panel A).
  • Pull both sides of the rubber band through the middle of the collar (panel B).
  • Then, while holding the collar, pull the rubber band down over the stopper and pinch the collar tightly around the neck of the bottle (panel C). Be sure to pull the collar down so that it is below the neck of the bottle. You may need to use channel locks to close the collar tightly and secure the stopper and rubber band in place. Special rubber bands are required for cold water sampling.
  • Invert the sample several times to mix the mercuric chloride thoroughly.
  • Dip each bottle in a bucket of clean fresh water up to the neck, dry, and place in storage crate. Bottles can be stored at room temperature but should be kept out of direct sunlight or high temperature. If cold storage is available, that is preferable, but samples should never be frozen.
  • The sample in panel D has the correct volume of headspace and a properly fit clip and rubber band.
  • Take duplicate samples as you see fit, e.g. 10% of your samples. Consider taking duplicates of all critical samples.

Sampling in estuarine or glaciated environment

Inorganic carbon sampling in glaciated coastal regions requires methodological variations from open-ocean best practices to ensure that suspended mineral particles do not compromise the instrumentation and/or bias DIC, TA, and pH measurements between sample collection and analysis [Sejr et al., 2011]. In these settings, the discrete seawater samples need to be filtered (replaceable 0.45 µm filter in a 47 mm polycarbonate In-Line filter) with a peristaltic pump straight from the Niskin bottles [see Bockmon and Dickson, 2014 for detailed method], into pre-cleaned 500 mL borosilicate bottle, and poisoned with mercuric chloride (HgCl2) [Dickson et al., 2007]. It is also advised to distinguish between carbonate and non-carbonate alkalinity in this environment by over-constraining the inorganic carbon system by measuring three of the required two variables [Byrne, 2014].

References

Byrne, R. H. (2014), Measuring Ocean Acidification: New Technology for a New Era of Ocean Chemistry, Environ. Sci. Technol., 48, 5352–5360, doi:10.1021/es405819p.

Dickson, A.G.; Sabine, C.L. and Christian, J.R. (eds) (2007) Guide to best practices for ocean CO2 measurement. Sidney, British Columbia, North Pacific Marine Science Organization, 191pp. (PICES Special Publication 3).

PMEL Carbon Program: https://www.pmel.noaa.gov/co2/files/dic_sample_technique_revised_9-5-2018.pdf

https://www.pmel.noaa.gov/co2/story/Laboratory+analysis+details

Sejr, M. K., D. Krause-Jensen, S. Rysgaard, L. L. Sørensen, P. B. Christensen, and R. N. Glud (2011), Air-sea flux of CO2 in arctic coastal waters influenced by glacial melt water and sea ice, Tellus B, 63(5), 815–822, doi:10.1111/j.1600-0889.2011.00540.x.

Overview of Best Practices is adapted from PMEL’s website.