Data Parameters and Units:
Biomass - date (MM/DD/YYYY), Local Time, GMT time (hh:mm), Latitude (decimal degrees), Longitude (decimal degrees), Station, Bottom Depth (meters), Estimated Two Depth (m), Volume Filtered (meters^3), Biomass (mg/meters^3). Chlorophyll- date (MM/DD/YYYY), Local Time, GMT time (hh:mm), Latitude (decimal degrees), Longitude (decimal degrees), Station, Depth (meters), Chlorophyll (mu g/L), Phaeopigments (mu g/L), Phaeo/Chl (mu g/L) [Whole Sample and Size Fraction greater than 3 mu m]. CTD-- NT: Scan, date (MM/DD/YYYY), GMT time (hh:mm), Latitude (decimal degrees), Longitude (decimal degrees), Depth (meters), Temperature (deg C), Potential Temperature (deg C), Salinity, Conductivity (S/m, mS/cm), Density (sigma-theta kg/m3), Pressure (dB), Oxygen (ml/l, mg/l, mu mol/kg, % saturation), Fluorescence (DOM mg/m^3, ECO-AFL/FL mg/m3), Turbidity (NTU), Beam Attenuation (1/meters), Beam Transmission (%), Average Sound Velocity (Chen-Millero m/s, Delgrosso m/s, Wilson m/s). Nutrients - date (MM/DD/YYYY), Local time, GMT time (hh:mm), Latitude (decimal degrees), Longitude (decimal degrees), Station, Depth (meters), Urea (mu M N), NO3 (mu M), NO2 (mu M), Silica (mu M), NH4 (mu M), PO4 (mu M), TDN Total Dissolved Nitrogen (mu M), TDP Total Dissolved Phosphorus (mu M). SIPPER - latitude deployed, longitude deployed, latitude recovered, longitude recovered, depth (m), volume filtered (m-3), abundance per m^3 of: Chaetognath, Crustacean, Crustacean copepod Evadne, Crustacean cladoceran Penilia, Crustacean copepod, Crustacean calanoid copepod, Crustacean copepod Copilia, Crustacean copepod Macrostella, Crustacean copepod Macrostella Tricho, Crustacean copepod nauplii, Crustacean copepod Oithona, Crustacean copepod Poecilostomatoid, Crustacean copepod Sapphirina, Crustacean eumalcostracan euphausiid, Crustacean eumalcostracan Lucifer, Crustacean ostracods, Crustacean phyllosome, detritus, detritus blob, detritus molts, detritus snow, echinoderm, echinoderm bipinnaria, echinoderm plutei, eggmass, fish, gelatinous zooplankton, gelatinous ctenophore, gelatinous hydromedusae, gelatinous siphonophore, gelatinous tunicate doliolid, gelatinous tunicate pyrosome, gelatinous tunicate salp, lancelet, larvae, larvae doliolid, larvae polychaete, larvae tornaria, larvae veliger, mollusc, mollusc Atlanta, mollusc heteropod, mollusc pteropod, mollusc pteropod conical, mollusc pterpod gymnosome, mollusc pterpod pseudothecosomata, mollusc pteropod shell, mollusc squid, other, phytoplankton, phytoplankton diatom centric, phytoplankton diatom coil, phytoplankton strands, phytoplankton Trichodesmium, polychaete, protist, prostist Acantharia, protist Collozoum, protist electric, protist knobby, protist Noctiluca spp., Protist radiolarian, protist Thalassicola spp., protist unknown, temperature (degrees Celsius), Salinity, Density (kg/m^3), Fluorescence (mg/mg3), Fluorescence (volts), oxygen (mL/L), oxygen (umol/kg), transmissivity (% light), turbidity (NTU), CDOM Fluorescence (ppb QSD). Zooplankton abundance - date sampled (MM/DD/YYYY), station ID, Latitude (degrees N), Longitude (degrees W), local time, GMT Time (HH:MM), Volume filtered, bottom depth (m), estimated sampling depth (m), split, abundance (#/m3) of: noncalanoid copepods, calanoid copepods, other taxa.
Methods:
CTD: Environmental data were collected aboard the R/V Weatherbird II and the R/V Bellows. A CTD-rosette was deployed between the surface (2 m) and up to a maximum depth ranging from 5 m to about 1,450 m, depending on the bottom depth of the station. Sensors mounted on the rosette included: SBE 25 CTD, SBE 43 DO, Biospherical PAR sensor, WET Labs CDOM (FLCDRTD), Wet Labs fluorometer and turbidity sensors (FLNTURTD), and WET Labs C-Star transmissometer. The CTD was deployed to ~ 2 m depth, the power turned on, and allowed to sit a few minutes until the pumps turned on and sensors stabilized and equilibrated. The CTD was then lowered at a rate of 30 m/min for the first 100 m, and thereafter at 60 m/min. On the downcast, the depths of the fluorescence maxima, top of the thermocline, base of the thermocline, and other features of interest were noted on the CTD log sheet. Niskin bottles were fired at these depths and other pre-determined depths on the upcast. Water samples were collected during more than one CTD cast at some stations where water demand was high or the depth was >200 m. The Weatherbird II rosette held 12-12 L Niskin bottles, and the Bellows rosette held 12-5 L Niskin bottles. Gloves were worn to collect water samples. The order of water samples drawn from Niskin bottles was: gases, nutrients, chlorophyll, pigments (HPLC), CDOM, other. Raw data were saved as hex files and converted to 1 m bin averaged data using the Sea-Bird Electronics Inc. Data Processing Software v.7.23.2. Chlorophyll: Chlorophyll samples were collected in 2 L amber bottles using tygon tubing without mesh for depths between 2 to 500 m. Gloves were worn for sample collection and bottles and caps rinsed three times before sample collection. The sample bottles were immediately processed after collection. The collection bottle was gently swirled and 500 mL to 2 L was filtered under low vacuum onto a 25 mm GF/F filter. The filter was then placed in a foil packet and labeled, and frozen at -20 C. Some field samples were size-fractionated. In the lab, the filter was placed in a 13 x 100 mm borosilicate tube and then 7 mL of 90% acetone was added. The tube was covered with parafilm and placed in a dark box in a -20 C freezer for 24 hours. After the 24 hr extraction, samples were allowed to come to room temperature (10-15 min) in the dark. Chlorophyll a concentrations were determined using a Turner fluorometer 10-A before and after acidification following the methods described in Holm-Hansen et al. (1965). The fluorometer was calibrated every six months using a Sigma-Aldrich chlorophyll a standard. Reference: Holm-Hansen, O., C.J. Lorenzen, R.W. Holmes, and J.D. Strickland. 1965. Fluorometric determination of chlorophyll. J. Cons. Cons. Int. Explor. Mer 30: 3-15. Nutrients: Nutrients were collected in acid cleaned wide mouth HDPE 30 mL bottles. Gloves are worn while collecting samples. Bottles and caps were rinsed three times, filled up to the shoulder of the container, and then frozen upright at -20 C. Samples were analyzed for nitrate (NO3 µM), nitrite (NO2 µM), phosphate (PO4 µM), silica (SiO2 µM), and ammonium (NH4 µM) by R. Masserini, A. Yunker, and K. Fanning (USF) following recommendations of Gordon et al. (2000) for the WOCE WHP project. The analytical system employed is a five-channel Technicon Autoanalyzer II upgraded with new heating baths, proportional pumps, colorimeters, improved optics, and an analog to digital conversion system (New Analyzer Program v. 2.40 by Labtronics, Inc.) Silica was determined by forming the heteropoly acid of dissolved orthosilicic acid and ammonium molybdate, reducing it with stannous chloride, and then measuring its optical transmittance. Phosphate was determined by creating the phosphomolybdate heteropoly acid in much the same way as with the silica method. However, its reducing agent is dihydrazine sulfate, after which its transmittance is measured. A heating bath is required to maximize the color yield. Nitrite was determined by the Bendschneider and Robinson (1952) technique, in which nitrite is reacted with sulfanilamide (SAN) to form a diazotized derivative that is then reacted with a substituted ethylenediamine compound (NED) to form a rose pink azo dye, which is measured colorimetrically. Nitrate was determined by difference after a separate aliquot of a sample is passed through a Cd reduction column to covert its nitrate to nitrite, followed by measurement of the "augmented" nitrite concentration using the same method as in the nitrite analysis. In the analytical ammonia method, ammonium reacts with alkaline phenol and hypochlorite to form indophenolblue. Sodium nitroferricyanide intensifies the blue color formed, which is then measured in a colorimeter on our nutrient-analyzer. Precipitation of calcium and magnesium hydroxides is eliminated by the addition of sodium citrate complexing reagent. A heating bath is required. The USF/CMS version of this technique is based on modifications of published methods such as the article by F. Koroleff in Grasshoff (1976). These modifications were made at Alpkem (now Astoria-Pacific International, Inc.) and at L. Gordon's nutrient laboratory at Oregon State University. Detection Limits (micromolar) for Nutrient Analyses: Nitrate + Nitrite = 0.22, Nitrite = 0.02, Silicic Acid = 0.29, Ammonium = 0.38, Phosphate = 0.09. Reference: Gordon L.I.; Jennings Jr, J.C.; Ross, A.A.; Krest, J.M. 2000. A Suggested Protocol For Continuous Flow Automated Analysis of Seawater Nutrients. WOCE Operation Manual, WHP Office Rept 90-1, WOCE Rept 77 No 68/91, 1-52; revised by Ross, A.A. Urea Measurements. To determine urea concentrations, seawater samples were collected from Niskin bottles using tygon tubing with 153 µm mesh filter attached to remove large zooplankton. Gloves were worn during sample collection. Sample waters were passed through pre-combusted (450° C, 2.5 hrs) 25 mm Whatman GF/F filters (nominal pore size 0.7 µm) under low vacuum pressure and duplicate 10 mL aliquots of filtrate were poured into HDPE plastic scintillation vials and frozen (-20° C) until processed. In the lab, 10 mL of sample was pipetted into glass tubes containing 1.1 g NaCl. Tube racks were placed in a cool pan of water and 1.4 mL of the sulfuric acid reagent was added. After mixing the sample, 0.3 mL of the diacetylmonoxime reagent was added to the sample tube and vortexed until all NaCl was dissolved. The tube racks were then placed in an oven and heated at 75° C for two hours. Samples were read at an optical density of 520 nm on a Beckman DU720 Spectrophotometer. Detection Limit = 0.05 micromolar. Reference: Koroleff, F. 1976. A modified manual method for the determination of urea in seawater using diacetylmonoxime reagent. Estuarine, Coastal and Shelf Science 34 (5): 429-438. Total Dissolved Nitrogen Samples Seawater samples were collected in acid cleaned wide mouth HDPE 30 mL bottles and analyzed following methods in Valderrama (1981). Gloves were worn during sample collection. Total dissolved nitrogen (TDN) concentrations were determined by the oxidation of dissolved organic nitrogen (DON) in filtered samples to nitrate. Samples are treated with potassium persulfate, boric acid, and sodium hydroxide and then autoclaved for 30 minutes at 121°C and 15 lb/in^2 pressure to convert DON, ammonium, and nitrite to nitrate. The nitrate concentrations of the oxidized samples, determined colorimeterically following the recommendations of Gordon et al. (2000), are equal to the TDN. DON is derived by subtracting the total inorganic nitrogen (TIN) value, determined seperately, from the TDN value. Reference: Valderrama, J. C. 1981. The simultaneous analysis of total nitrogen and total phosphorus in natural waters. Mar. Chem. 16: 109-122. Total Dissolved Phosphorus Measurements. To determine total dissolved phosphorus (TDP), seawater samples were collected from Niskin bottles using tygon tubing with 153 µm mesh filter attached to remove large zooplankton. Gloves were worn during sample collection. Sample waters were passed through pre-combusted (450° C, 2.5 hrs) 25 mm Whatman GF/F filters (nominal pore size 0.7 µm) under low vacuum pressure and duplicate 10 mL aliquots of filtrate were poured into fired scintillation vials (450° C, 2.5 hrs). Samples were then treated with 0.2 mL of 0.17 M MgSO4 and frozen (-20° C) until processed. In the lab, scintillation vials were placed in a drying oven at 95° C. Upon desiccation, the scintillation vials were fired in a muffle furnace at 450° C, for 2.5 hours. The samples were than hydrolyzed with 3 mL of 0.75 M HCl and heated for 20 minutes. An additional 7 mL of distilled water was added to the vial and heated for another 10 minutes. The samples were then transferred into test tubes, treated with 1 mL of mixed reagent and read at an optical density of 885 nm on a Beckman DU720 Spectrophotometer. Detection Limit = 0.05 micromolar. Reference: Solarzano, L. & J.H. Sharp. 1980. Determination of total dissolved phosphorus and particulate phosphate in natural waters. Limnol. Oceanogr. 25: 754-758. Bongo nets: Sample Collection. Bongo nets (60 cm diameter, 505 µm mesh nets) were towed obliquely through the water column from 200 meters to the surface, while the ship was underway at 1 - 2 knots keeping an approximate 45 degree wire angle. At stations shallower than 200 m, the maximum sampling depth was targeted to be 5 m above the bottom. The wire was let out at 50 m/min until the maximum depth was reached, held at depth for 30 seconds, and then retrieved at a constant rate (20 m/min). Flow meters in each net were read before and after each net deployment. The sample in the right cod end was preserved in buffered 10% formalin. The sample in the left cod end was concentrated onto a 200 μm sieve using salt water. The sample was photographed and then scooped into Falcon tubes or combusted 30 mL glass vials with screw caps for biomass analyses. The biomass tubes were placed into a labeled freezer bag and frozen at -20° C. Bongo nets: Zooplankton Biomass Analyses. Zooplankton samples were removed from the freezer and allowed to thaw to room temperature. The samples were then placed into pre-weighed aluminum boats and dried at 60º C for about one week. Once dried, the samples were removed from the oven, allowed to cool to room temperature, and then reweighed. Zooplankton dry weight (mg) = (zooplankton + boat weight) – weight of boat. Zooplankton dry weight biomass (mg/m3) was calculated as the weight of zooplankton (mg) divided by the volume of water filtered (m3) by the bongo net for each specific tow. Bongo nets: Zooplankton Identification and Abundance Analyses. Large Taxa Identification. In the laboratory, the entire preserved sample was poured onto a 64 µm mesh sieve and rinsed with filtered seawater; it was then poured into a large clear pyrex dish for sorting. Large (easily detectable by eye) non-copepod taxa (e.g. sergestids, myctophids, mysids, heteropods, pteropods, large chaetognaths, hyperids, euphausiids, etc.) were rinsed with filtered seawater and removed for later enumeration and identification. These taxa were identified to group or species (see Gulf Mexico taxa Sept 2012.doc) and then stored in glass vials with 4% sodium borate buffered fomaldehyde in filtered seawater. Some taxa were categorized further to life stage (e.g. (fish larvae: preflexion, flexion and post flexion). Copepods and Other Small Taxa Identification. To obtain abundances of copepods and other small taxa, the preserved sample with large taxa removed was again poured onto a 64 µm mesh sieve and rinsed with filtered seawater into a Folsom plankton splitter. It was then split to between 1/8 and 1/1024 of the whole sample, depending on the abundance of calanoid copepods. The goal was to obtain approximately 100 calanoid copepods. This split was then sorted using a dissecting microscope (15 - 60 X). Calanoids (e.g. Eucalanidae, Metridiidae, Aetidiidae, etc.) and non-calanoids (e.g. Corycaeus, Lubbockia, etc.) were removed, identified, enumerated and then stored in glass vials with 4% sodium borate buffered fomaldehyde in filtered seawater. Calanoids were identified to the lowest possible taxon (see Identification Methods and Notes below) and staged. In some cases (e.g. Paracalanus spp., Clausocalanus furcatus, Ctenocalanus vanus, etc.), identification was confirmed under a compound microscope. Non-calanoids were identified to genera (e.g. Oncaea, Oithona, Corycaeus, etc.), but not staged. If damage to the copepod prevented identification or staging, the individual was labeled “damaged” and placed in the lowest category possible. Small non-copepod taxa (e.g. ostracods, cladocerans, larvaceans, foraminiferas, echinoderm larvae, etc.) also were removed from the split and identified to group or species, enumerated, and stored in glass vials. Calculations to Determine Abundances (number per m3). All copepods (calanoids and non-calanoids) were identified from the sample split (1/8 to 1/1024). All other non-copepod taxa were removed from the whole and/or a split. To obtain abundances for animals removed from the whole sample, prior to splitting, the raw value was adjusted for volume filtered (raw count ÷ volume filtered). To obtain abundances for animals from the split, the raw values were calculated as follows: ([raw count * split correction] ÷ volume filtered). Split correction is the inverse of the split (e.g. 1/8 = 8). On two occasions (PCB03 [December 2010] and PCB04 [December 2010]) the initial lab sample was previously split at sea to 1/2; therefore these samples were adjusted for both at sea split and volume filtered. An overall abundance (#/m3) for each non-copepod taxa was calculated by combining the removed from whole abundance with the split abundance. Due to high volumes of phytoplankton in the February 2011 samples, some samples had an intermediate split enumerated for certain large taxa (i.e. organisms scarce enough to potentially not make the copepod split, yet too abundant to fully remove from the whole with the phytoplankton). SIPPER (Shadowed Image Particle Profiling and Evaluation Recorder) Deployments: The The Shadowed Image Particle Profiling and Evaluation Recorder (SIPPER) is a towed, battery-powered, in-situ particle imaging system developed for studying the distribution and abundance of zooplankton and suspended particles in the water column (Samson et al., 2001, Remsen et al., 2004). It utilizes a collimated LED light source and a high speed line scan camera to continuously image particles as they pass through a 9.6 cm × 9.6 cm (92 cm2) sampling aperture as the system is towed through the water. The linescan camera images the 9.6 cm depth of field onto a single 4096 pixel detector array, that is scanned at rates up to 30 kHZ. The resulting two-dimensional image is a brightfield record in which the clear water background appears white and suspended particles and plankton appear as silhouettes and outlines in 3-bit grayscale. The effective optical resolution of the SIPPER is ~70 µm across a 9.6 cm depth of field and allows for the capability to distinguish between major plankton groups ranging in size from 0.5 mm to 5 cm or more in total length. SIPPER was deployed at 2 – 3 knots between 2 m (surface) and 5 to 20 m above the bottom or to a maximum of depth of 300 m, depending on the depth of the station. The system was towed in an oblique fashion to allow for 1 to 3 minutes of imaging at each one-meter depth interval. Environmental data was collected concurrently with imaging. Environmental data from external commercial off the shelf sensors included salinity, temperature, depth, dissolved oxygen, chlorophyll fluorescence, and transmissivity. SIPPER data are transferred to a personal computer upon retrieval and processed using the Plankton Image Classification and Extraction Software (PICES) and managed with the PICES Commander image database system. SIPPER data are classified using multi-class support vector machine classifiers (Luo et al. 2004, 2005) based training libraries built by experts familiar with Gulf of Mexico plankton and particle groups. Classification performance is determined by manually validating random subsets of images from select deployments and comparing those results against the results from the automated classifier. These random subset classifications also allow for the customization of classifiers for given cruises, regions, and time periods, such that classifiers only contain image groups known to occur for each deployment. References: Luo, T. et al. 2004. Recognizing Plankton Images from the Shadow Image Particle Profiling Evaluation Recorder. IEEE Transactions on Systems Man and Cybernetics Part B – Cybernetics 34 (4): 1753-1762. Luo, T. et al. 2005. Active Learning to Recognize Multiple Types of Plankton. Journal of Machine Learning Research 6: 589 – 613. Remsen, A., S. Samson, T. Hopkins. 2004. What you see is not what you catch: A comparison of concurrently collected net, optical plankton counter (OPC), and Shadowed Image Particle Profiling Evaluation Recorder (SIPPER) data from the northeast Gulf of Mexico. Deep Sea Research I 51(1):129-151. Samson, S., T. Hopkins, A. Remsen, L. Langebrake, T. Sutton, and J. Patten. 2001. A system for high resolution zooplankton imaging. IEEE Journal of Oceanic Engineering 26:671-676.
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