ESCI 322 - Oceanography Laboratory

Laboratory Manual

Prepared by David Shull Department of Environmental Sciences Huxley College of the Environment Western Washington University Bellingham, WA 98225 [email protected] Last update: September 20, 2011 ECSI 322 – Oceanography Laboratory - Manual

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ESCI 322 - Introduction to Oceanography Laboratory Course Syllabus Instructor: David Shull Office: ES 445, ext. 3690 Email: [email protected] OBJECTIVES: 1. Acquire first-hand experience with oceanographic research methods 2. Become familiar with marine organisms and coastal processes in Puget Sound 3. Learn field and laboratory techniques 4. Practice scientific writing (writing proficiency course - WP3) 5. Use oceanographic methods to address local marine environmental problems. COURSE OVERVIEW: We will use oceanographic methods to study ecosystem functions and environmental problems in the marine waters near Bellingham. Students will write reports addressing these issues. Many of our "labs" will take place in the field, either on a boat or at the Shannon Point Marine Center. EVALUATION GUIDELINES: Assignments: Students will be evaluated based on the completion of several lab reports, a few smaller lab assignments, and “pre-laboratory” assignments. Each assignment will be typed, although figures and graphs may be drawn by hand. The reports will be typed and will follow the standard scientific format; abstract, introduction, methods, results, discussion, references. For some of the reports, a draft of a portion of the results section will be handed in first for evaluation and comments. Comments on the draft of the results section are intended to aid students in the completion of the final reports. Both the results-section draft and the final reports must be turned in on time to receive full credit. Late reports will receive a 10% grade reduction each day until the report is turned in. Details on the format of the major reports are given in the section of the syllabus entitled "Laboratory Report Format". Read this section carefully before you begin. The smaller assignments will be completed in question-and-answer format. All reports should be typed, double-spaced, and checked for correct grammar and spelling. You should read through the assignment, make notes, and think through the organization of all your responses before writing. Pre-laboratory assignments needn’t be typed but must be handed in at the beginning of each laboratory period to receive credit. Contributions of assignments to final grade: Full lab reports Draft Reports Pre-lab assignments

80% 10% 10%

Approximate grading scale: 93-100 A 90-92 A88-89 B+ 73-77 C 69-72C67-68 D+

80-82 B57-60 D-

83-87 B 61-66 D

ECSI 322 – Oceanography Laboratory - Manual

78-79 C+ 0-56 F

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Policy for late assignments and reports: Final reports will lose 10% of the final grade for each late day until turned in. (Reports received after 5 PM will be picked up the next day.) Pre-labs and draft reports will not be accepted late. Lab schedule: Date 27-Sep 4-Oct 11-Oct 18-Oct 25-Oct 1-Nov 8-Nov 15-Nov 22-Nov 29-Nov

Topic Waves Habitat utilization by rockfish I Habitat utilization by rockfish II Water column profiles in Bellingham Bay Chlorophyll and phytoplankton biomass Nutrients in Seawater I Nutrients in Seawater II Measuring phytoplankton growth and grazing I Measuring phytoplankton growth and grazing II No class, final report due

Location ES 60 SPMC* WWU SPMC* SPMC ES 331 ES 331 SPMC SPMC WWU

11-Oct 18-Oct 25-Oct

Final Report 4-Oct 18-Oct

+ROV

1-Nov

+CTD

8-Nov

22-Nov

Draft

Zoea required

29-Nov

*On these days we will be outside for part or all of the lab session. Dress for cold, rain and wind

LAB REPORT FORMAT A laboratory report is a document in the form of a scientific paper. Scientific papers are the means by which scientists communicate their research findings to one another. Writing up and publishing research results is just as important as conducting research in the first place, for if results are not made available to others, they are of little value. For ease of communication, there is a generally accepted format for writing up scientific data that we will follow in this course. Mastery of scientific writing skills is a vital component of becoming a scientist. Scientific papers have 7 sections: Title, Abstract, Introduction, Materials and Methods, Results, Discussion, and References. These headings should be placed at the beginning of each section in your report (except “Title”). Brief descriptions of the seven sections follow. Title The title should be a self-contained explanation of the information presented in your paper. It must contain enough detail to be informative, without being so long as to be incomprehensible. Avoid vagueness at all cost. Too short: Vertical profiling Too vague: Oceanography laboratory report #1 Too long: A student investigation of the effects of the Nooksack River on the vertical and horizontal distribution of temperature, salinity, density, nutrients, and dissolved oxygen in Bellingham, Bay, WA, in November, 2010. Abstract The abstract is a short one- to two-paragraph essay that summarizes the major findings of the paper. The abstract is important because it may be the only part of the paper someone will read. ECSI 322 – Oceanography Laboratory - Manual

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If the abstract is interesting, concise, well-written, and accurately summarizes the content of your work, it could motivate someone with an interest in the topic to read the rest of the paper. The abstract must not include ideas or information that is not included in the rest of the paper. It should briefly discuss the motivation for the study, the major results, and inferences drawn from those findings. If the development of new methods is an important part of the paper, they should be described in the abstract as well. References are not typically cited in the abstract. Introduction The introduction sets the stage for the presentation of your research results and their interpretation. It must include some background information, to bring the reader up to speed on the general issues, some specifics, to acquaint the reader with your particular investigation, and the questions or hypotheses that you will be addressing with the data. Effective introductions are usually short (several paragraphs). Background information: What is the general problem that is being studied? What is your specific approach to that problem? If there is relevant background literature (other important scientific papers that set the stage for your work), this is the place to briefly summarize their findings and importance. However, the introduction should not be a literature review. Specifics: This section will vary depending on the type of research you are presenting. In an environmental study, you should let the reader know where and when you were working, and what the environment was like in a general way. If you were presenting the results of an experiment with organisms, you should describe the species used and the general approach. Try to develop a logical flow from the “big picture” of background information to the specifics of the system you studied. Research questions: End your introduction with a concise summary of the research questions, hypotheses or goals. You will come back to these in the concluding paragraph of your discussion. Materials and Methods The materials and methods section describes how, when, where and what you did. Describe the procedures, equipment, and experimental set-ups in enough detail that the work could be repeated by another scientist, but without extraneous detail. List the methods or procedures chronologically (i.e. in the order in which you did them). Be sure to include information about numbers of replicate treatments or observations, types of instruments and equipment used, etc. If statistical analyses were performed, state the statistical methods used and the data to which they were applied. Use the past tense. Methods that are already published can be referred to with a reference; only deviations from the published method need to be described in your paper. (“Nitrate was measured by the method of Parsons et al. (1984), except that reagent additions were scaled to a sample volume of 5 ml.”) You can reference lab handouts, but be sure they are cited in the reference section. You do not need to explain things that are not necessary for understanding or repeating the work (“The group was divided in half and group A went out in the boat first, then group B.”) You can use either passive voice (“Salinities were measured at 1-m intervals.”) or active voice (“We measured salinities at 1-m intervals.”), but I prefer active voice as it is generally more precise.

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Results The results section is the heart of the paper. This is where you tell the reader everything that you found: what, when, where. Interpretation of those data, however, is left for the discussion section. This allows the reader to formulate her or his own interpretation before reading yours. The results section consists of tables and graphs that summarize your data (not raw data), and text that describes and highlights the major features of those data. The results section text is often fairly short. Use the past tense here, as the observations were made in the past (e.g: Surface water salinity was lower at stations in northern Bellingham Bay near the mouth of the Nooksack River compared to stations further south.) Figures and Tables: Each figure (map, diagram, or graph) and table in a results section should have a “reason for being.” Don’t present data just because you collected it; present data only if it you refer to it in the text and it contributes to the story that you are telling in your paper. In general, figures are plotted with the independent variable on the x axis. Vertical profiles in oceanography have a special format in which the independent variable (depth is plotted on the yaxis. (We'll discuss this more in class.) Each table and figure should be readily understandable without reference to the text. Each should have a consecutive number (Fig. 1, 2, 3…); tables are numbered separately from figures. Finally, each figure and table should have a caption (sometimes called a ‘legend’) that concisely describes the content (e.g.: Fig. 2. Average rates of respiration (±1 s.d.) over time for the anemone Anthopleura elegantissima at 10ºC.) Text: The text of the results section should weave the data presented in figures and tables into a coherent story. Prepare the figures and tables first, and then write the text. Do not reiterate all the details of the data; rather, tell a story that describes the major features and any clear trends or patterns. Refer directly to the appropriate figures and tables, by number, in your text. The first figure referred to should be Fig. 1, the second Fig. 2, etc. Keep the writing simple and direct. Don't use the word Figure or Table as the subject of a sentence, e.g., Figure 1 shows the locations of the sampling sites, because this ruins the narrative style. Instead, tell the story and refer to figures in parentheses. Examples: Not good: The graph of temperature versus depth looks linear near the bottom. Still not so good: Fig. 1 shows that temperature was constant with depth near the bottom. Better: Temperature was constant throughout the bottom 5 m of the water column (Fig. 1). Incorrect: A paired t-test showed that the respiration measurements were significantly different at the 95% confidence level. Good: Growth rates in the anemones were higher at 12ºC than at 10ºC (t-test, t[8]=2.9, p = 0.02). Discussion In the discussion you interpret your results: tell the reader what they mean and why they are important. In this section you should answer “why?” and “how?” questions about your data. For example, why was the temperature different at the top relative to the bottom of the water column? Why did the respiration rate of the anemones increase with temperature? Here reader should discover the answers to your original questions or hypotheses as set forth in the introduction. The discussion section is also where you compare your findings to those of others, as reported in the scientific literature. You may also want or need to discuss short-comings in

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your methods, or the need for further testing. This latter should not, however, be the main focus of the section. The discussion section should be a general synthesis of your findings and their importance. Do not restate your results. The key word is interpretation. This section is usually the hardest to write; think it through carefully and prepare an outline before you begin. One effective technique is to start the section with your strongest or most important finding. References This section is an alphabetical listing, by first author’s last name, of the references cited in your paper. There are two ways to cite a paper in your text: Several other species of anemone are known to have respiration rates that increase with temperature (Matthews, 1993; Smith and Wesson, 1998). Our findings of lower respiration rate at lower temperatures agreed with those of Matthews (1993). If there are more than two authors, use the term et al. (an abbreviation of et alias, “and others”) after the name of the first author: Michaels et al. (1994) or (Michaels et al., 1994). Journals have their own specified format for listing references, which should be followed when submitting a paper to that journal. For our purposes, you can use the format below. Journal article: Name(s) of author(s). Date. Title of article. Title of journal (may be abbreviated). Vol #: pages. Example: Lenington, S. 1979. Effect of holy water on the growth of radish plants. Psychological Reports 45: 381-382. Book: Name(s) of authors. Date. Title of book. Publisher, Location (city) of publisher, # of pages in entire book. Example: Povinelli, D. J. 2000. Folk Physics for Apes: The Chimpanzee's Theory of How the World Works. Oxford University Press, New York, 391 pp. Chapter from a book in which each chapter has a different author and the book has an editor: Name(s) of authors. Date. Title of chapter. In: editor(s), Book Title. Publisher, Location (city) of publisher, pages. Example: Jumars, P. A. 1993. Gourmands of mud: Diet selection in marine deposit feeders. In: R.N. Hughes (Ed.), Mechanisms of Diet Choice. Blackwell Scientific Publishers, Oxford. pp. 124-156.

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ESCI 322 Lab Assignment : Waves and Coastal Processes During this laboratory session we will study some of the properties of waves. We will create our own experiments in wave tanks, altering variables known to affect wave properties (see below), to explore the mathematical relationships among those properties. We will also set up an experiment to simulate the effects of waves on coastlines. Terminology: -Wave height (H) is the vertical change in height between the wave crest and the wave trough. -Wave amplitude (A) is one-half the wave height. -Wavelength (L) is the distance between two successive peaks or troughs. -Steepness is wave height divided by wavelength (H/L) (note this is not the same as the slope between a wave crest and its adjacent trough). -Wave period (T) is the time it takes for two successive peaks (or troughs) to pass a fixed point. -Wave frequency (f) is the number of peaks (or troughs) which pass a fixed point per second. Review of wave relationships: Wave period is the inverse of wave frequency: T = 1/f Wave speed (termed celerity) can be related to wavelength and period according to the general formula: C = L/T. These formulas hold for all waves. For gravity waves at the water surface, the following equation can be used to calculate wave speed from wavelength and water depth. 1

 gL  2d   2 C   tanh   ,  L   2 where d is water depth (below mean surface level), and g is the gravitational acceleration (9.81 m/sec2). We will test this theoretical model in part two of our lab assignment. This equation reduces to simpler forms for deep-water and shallow-water waves. Deep-water waves: water depth (d) is greater than L/2 and C = gL 2 (m/s). Since g is a

constant, this formula reduces to C= 1.25 L (m/s). Note that L, the wavelength, is the only variable affecting wave speed for deep water waves. Since L, T and C are related, the equation for deep water waves can be rewritten as C = 1.56T (m/s). Shallow-water waves: water depth (d) is less than L/20 and C = gd = 3.13 d (m/s). Note that d, the water depth, is the only variable affecting wave speed for shallow-water waves. Intermediate waves: if water depth is < L/2 and >L/20, the more complex formula given above must be used to calculate wave speed. Affects of bottom topography: Refraction: Because wave speed for shallow-water waves is a positive function of water depth, waves slow as they approach the shoreline. Wave period remains constant so that a decrease in wave speed reduces wavelength. Parallel-crested waves approaching the shoreline at an angle, therefore, will refract, bending to become more parallel to the shore before they break. Bottom topography around bays and headlands will result in refraction patterns causing variation in the spatial distribution of wave energy, sediment erosion, and sediment deposition along the shore.

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Breaking waves: As waves approach the shoreline, steepness increases. Theoretically, waves become unstable and should break when the steepness (H:L) reaches 1:7. In practice, this ratio rarely exceeds 1:12 due to other sources of instability. Bottom topography also affects breaking so that breaking occurs when H/d equals about 0.8, regardless of H:L. Overview of lab procedures Large wave tank 1: Prepare a coastline in the large wave tank and measure its geometry 2: Turn on the wave generator in the large tank Narrow wave channel 3: Trace the beach profile in the narrow wave channel and measure the water depth 4: Turn on the wave generator and measure the wavelength and speed of waves 5: Change the wave period, height, and water depth and repeat the measurements 6: Measure the change in the beach profile 7: Measure the wavelength and water depth at which waves break Large wave tank 8: Return to the large wave tank, measure the coastline and assess the effects of waves Laboratory wave exercises Effects of waves on coastal processes We will conduct these experiments in the large wave tank. Begin by creating a coastline. Use shovels and other implements to create a coast at an angle to the oncoming waves. We will then be able to observe longshore current and longshore sediment transport. Pair up with a business partner and choose among the following jobs: Real-estate developer, marina developer/operator, park ranger, waste-water treatment plant operator, shipping company owner. Divide the coastline into equal allotments and develop it to suit your business needs. Use the materials in the tank to simulate the marina, homes, breakwaters, etc. Use the hose to simulate the sewer outfall. Make a careful drawing of the shoreline. Measure the distance of the shoreline from the edge of the tank at different locations. We will use these measurements later to identify areas of sediment erosion and deposition. Now, turn on the wave machine. Adjust the wave period by turning the control knob. Adjust the wave height by turning the knob on the wall to the right of the tank. Assignment – answer the underlined questions 1: Make a detailed drawing of the beach before turning on the wave tank Observe the directions of incoming waves relative to the shoreline at different distances from the shore. Do the waves refract as they approach the shore? Predictions 2: At which locations along the shoreline would you expect wave energy to be highest? 3: Where would it be the lowest? 4: Where would you expect rates of sediment erosion to be highest? 5: Where would you expect the rates to be lowest?

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Place a floating object in the tank near the shoreline. Can you observe longshore transport? Can you observe areas of erosion or deposition? Let the wave tank run while we move on to experiment two. After 1 h, repeat the shoreline measurements in the large tank and identify regions of erosion, deposition, and areas that are stable. Attempt to fix any problems with your property by dredging, adding a breakwater or groin, by beach nourishment, or by beach armoring. 6: Let the tank run for another hour and then repeat your drawing and observations. Results 7: Which of the development projects appear to be a success? 8: Which appear to be failures? 9: Why? Relationships between C, L, T, f, H, and d: These experiments will be conducted in the narrow wave channel. Turn on the wave generator and adjust the wave frequency by turning the small knob on the control box. Adjust the wave height by opening or closing the valve on the compressed air tank. Assignment Measure C, L, T, f, H, or d for waves of different heights and frequencies. Use these measurements to test the following relationships for transitional waves: Wave speed: Properties of breaking waves:

 gL  2d tanh  C    L  2

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 2   

H/L = 1/12 (= 0.083) H/D = 0.8

Measuring wave velocities and wave lengths is more difficult than it seems. We will use simple submerged pressure sensors attached to an oscilloscope to measure wave period (T) and speed (C). We will then calculate wavelength from the relationship L = CT. An oscilloscope displays a graph of an electrical signal. It shows how electrical signals vary over time; the vertical axis represents voltage and the horizontal axis represents time. As a wave passes over the submerged pressure sensor, the pressure increases according to the formula P=ρgh, where P is pressure, ρ is fluid density, and h is the height of the water above the sensor. The pressure sensors send a signal (a change in voltage) which is recorded on the oscilloscope. 10: Make two plots: (a) Measured wave velocity vs. water depth, and (b) measured wave velocity versus wavelength. Plot the measured wave velocities and the velocities predicted by the formula on the same graph. 11: How do your measurements compare with the established formula? ECSI 322 – Oceanography Laboratory - Manual

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Use a marker to draw the beach profile on the glass of the narrow wave channel. Draw this beach profile in your notes. Observe how the beach profile changes under different wave fields. Draw the new beach profile. 12: How do waves of different heights affect the beach profile? With a ruler, measure wave height and water depth at the point where waves break at the artificial beach for waves with different wavelengths. 13. Which ratio controls wave breaking (H/L, H/D)? Assignment Use your measurements of the coastline to help you draw it before and after wave exposure. Use your drawings and coastline measurements to identify areas of erosion and deposition in the big tank. Plot wave velocity in the wave channel versus water depth and wave length. Use these data and beach profile drawings to answer the questions from part two. (Just answer all the underlined questions in parts one and two. This is not a formal report.) Wave velocity data sheet Water depth

Wave period

Wave velocity

ECSI 322 – Oceanography Laboratory - Manual

Wavelength

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Assignment summary 1: Patterns of erosion and deposition due to wave action (large wave tank) - Draw developed shoreline before and after period of wave inundation - State how you expected the beach to change - Describe how it actually changed - Which development projects worked and which did not? Why? 2: Testing the wave velocity formula (wave channel) - Make two plots: Measured wave velocity vs. water depth, and measured wave velocity versus wavelength. Plot the measured wave velocities and the velocities predicted by the formula on the same graph. - Discuss whether the predicted wave velocities matched the measured velocities, and try to explain any differences you observe. 3: Effects of breaking waves (wave channel) - Draw two beach profiles (before and after wave inundation) - Calculate the ratios H/L and H/D at the location where the waves broke - Compare the measured ratios with the critical ratios for wave breaking - Which controls wave breaking in the wave channel?

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Pre-laboratory Report 1, Rockfish Habitat Utilization Name: _______________________________ Read the description of this week’s laboratory assignment and answer the following questions. You are watching video of the bottom collected by an ROV. In the center of the images you are viewing, two glowing points from laser beams are visible. The laser beams on the ROV are 10 cm apart. You measure the distance between the laser dots on the video screen and determine that the average distance between them on the video screen is 3 cm. The width of the image on the screen is 15 cm. You count six fish as the ROV traverses 50 m of bottom. Questions: What is the actual width of the video quadrat in cm? What is the area of the transect in m2? What is the fish density (number per square meter) in the transect?

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ESCI 322 Lab Report 1: Benthic habitat utilization Demersal fish and other bottom-dwelling organisms are often associated with particular types of bottom habitat. These organisms might seek out these habitat types because they provide food or refuge from predators. The larval stages of these organisms could be passively deposited in certain habitats or they might actively choose a particular bottom type for settlement. Alternatively, adults might migrate to these areas. Regardless of the reason behind these associations, habitat is clearly important to organisms and proper management of marine resources requires that appropriate habitats are preserved along with species of interest. Habitat preservation seems particularly important for rockfish in Puget Sound and the Strait of Georgia. Rockfish populations have been declining in recent decades and many advocate for the establishment of marine protected areas in Puget Sound to protect rockfish and their habitat. But, what kinds of habitat should be protected? The purpose of this week’s and next week’s labs is to document patterns of habitat utilization by rockfish in Whatcom County using a remotely operated vehicle, or ROV. The ROV is essentially a submersible video camera with propellers. It is tethered to the surface and controlled using a joy stick. We will deploy the ROV at several locations along the southeastern shore of Lummi Island where there exists a gradient in bottom type. We will deploy the ROV in a manner similar to that depicted in figure 1. Figure 1. ROV deployment and video collection plan (from Auster et al. 1997). Our survey will have two components. For the first, we will determine the bottom slope using the R/V Zoea’s GPS to determine our horizontal position (distance offshore) at several locations and the depth sounder to determine the water depth at the same locations. We will then be able to calculate bottom slope from the plot of bottom depth versus distance offshore. For the second component, we will drop the ROV at the deepest part of the sloping bottom and fly the ROV upslope, toward shore. We will direct the video camera toward the bottom and record fish and other organisms within the field of view of the camera. The ROV is equipped with two parallel lasers, 10 cm apart, which serve as a horizontal distance scale for the video. The lasers are set at a known angle from the horizontal so that the vertical distance scale can also be determined. To simplify the scale conversions, the camera tilt should be adjusted so that the laser dots are centered within the video field of view (Fig. 2). If L is the distance between the two lasers, and l is the distance between the two laser dots on the video screen, and a is the width of the video quadrat, then

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the actual width of the video, A, is equal to (L/l)a. If θ is the angle between the laser and the horizontal, and b is the height of the video frame, then the actual height of the frame, B, equals (L/l)b/sin(θ). If we know the length of the transect and its width, we can calculate the area surveyed. Rockfish density is the number of rockfish per unit area. We’ll determine transect width (A) from the distance between lasers. We’ll calculate the transect length from the change in water depth observed during the video transect and the bottom slope.

camera B Ll

θ

laser A H θ

H’’

Figure 2. Relationships between quadrat height and width and the laser beam angle θ. Analyzing the video data – week two During the second week we will analyze the video data. We’ll calculate the width and length of each transect. And, we’ll count and identify all of the rockfish we record on each transect. We’ll also characterize the bottom type of each transect (e.g., sand, boulders, gravel). Draft report For your draft report, you’ll calculate bottom slope from the GPS and water depth data. First, use the latitude and longitude from the GPS data to calculate the distance between each location where we took a water depth reading using the depth sounder. Then, plot the distance among locations versus water depth. Calculate the bottom slope of each transect (in degrees). Final report Your task is to address the following questions. What is the most important type of habitat for rockfish? What type of habitats should be protected as part of efforts to promote rockfish recovery? Determine the relationship between bottom slope and rockfish density (rockfish m-2). Assess how other aspects of bottom habitat might influence rockfish distribution. For example, did you observe spatial relationships between fish and particular features of the bottom habitat? Also determine the depth distribution of each rockfish species. Include a few references from the literature on rockfish and marine protected areas to bolster your report. Reference Auster, P,J., R. J. Malatesta and C. L. S. Donaldson. 1997. Distributional responses to smallscale habitat variability by early juvenile silver hake, Merluccius bilinearis. Environmental Biology of Fishes 50: 195–200.

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Pre-laboratory Report 2, Vertical profiles in Bellingham Bay Name: _______________________________ Read the description of this week’s laboratory assignment and access the Excel spreadsheet water_profiles.xls found on the course web site. Then, answer the following questions to be turned in at the beginning of the lab period. You do not need to type your answers to these questions. You may write your answers in the space available and turn in this sheet. 1: Water has a temperature of 14 °C and salinity of 34.5 %0. What is the density in kg m-3?

Density (kg m-3)

2: Use the equation of state to calculate the density of fresh water over the temperature range of 0 °C to 10 °C in 0.5 °C increments. Sketch a plot of temperature (x-axis) versus density (y-axis). According to your plot, at what temperature does fresh water reach maximum density?

Temperature of maximum density: _______

Temperature (°C)

3: Seawater has a density of 1.027 kg/l. What is the density of the seawater in t? ___________

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ESCI 322 Lab Report 2 Part 1, Vertical profiles in Bellingham Bay The Nooksack River enters Bellingham Bay at its northern end. It delivers approximately one billion cubic meters of freshwater to Bellingham Bay each year. This significantly reduces the salinity of the bay and affects the mean circulation in the bay as well. The Nooksack also delivers nutrients and other dissolved solutes and organic matter. Thus, it strongly affects the biology and chemistry of Bellingham Bay. Recently, regions of low dissolved oxygen (DO) concentrations have been observed in the center of the bay during late summer. A similar pattern of low oxygen (termed hypoxia) has been observed in other small embayments throughout Puget Sound. Furthermore, there are plans to double the size and output of the Post Point Wastewater Treatment Plant, which empties into Bellingham Bay, by 2014. How will this change in nutrient and freshwater input to Bellingham Bay affect its nutrient and oxygen concentrations? The objective of today's lab is threefold. First, we’ll examine how freshwater input affects the chemistry and physics of Bellingham Bay. Second, we’ll observe the distribution of dissolved oxygen in the water column and sediment and consider what processes control DO concentration. (Next week we’ll look at similar processes in the sediment.) Third, the data we collect will contribute to an ongoing monitoring program on water quality and dissolved oxygen in Bellingham Bay that I have been conducting with my classes. Important water column properties: Nutrients: The primary limiting nutrient in coastal marine systems is typically nitrogen, although phosphorus availability may limit productivity as well. Availability of silica can limit the productivity of diatoms, which have silica frustules (outer shells). Nitrogen occurs in several forms – ammonium (NH4+), nitrite (NO2-) and nitrate (NO3-). Nitrogen waste products are released into seawater as NH4+ or as a compound such as urea that is quickly converted to NH4+. Ammonium in seawater is oxidized to form NO2- which is then oxidized to NO3- by nitrifying bacteria. Phosphorus is found primarily as HPO42- in seawater. Its chemical form varies with seawater pH. Silica (silicic acid) is found mostly as Si(OH)4 at seawater pH. The productivity of coastal marine ecosystems is strongly dependent on concentrations of these nutrients. Light intensity: Primary production is also strongly affected by light intensity. Light interacts with algal pigments to drive photosynthesis; both the quality (spectrum, or color) and quantity of light are important in regulating this process. Secchi disk: A black and white disk is a low-tech way to measure the penetration of light into water. Named for Father Pietro Angelo Secchi, this primitive instrument has been used extensively in marine and aquatic systems for studying irradiance. Today, when the quantum sensor smashes against the side of the ship, the Secchi disk comes out. Lower the disk until it can no longer be seen, then raise it to the depth at which it just becomes visible again. Record this depth. You may have to repeat this several times to obtain a consistent Secchi depth. Spherical light sensor (4pi):This spherical light sensor measures the total amount of photosynthetically active radiation (PAR, radiation at wavelengths that can be used in photosynthesis). PAR wavelengths range from 400 to 700 nm for most photosynthetic organisms. Although light enters the water from above, it is scattered by water molecules so that,

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from an underwater vantage point, light comes from all directions. The spherical (4pi) sensor allows accurate measurement of this diffuse light field. Measuring phytoplankton pigments: The CTD has a fluorometer that measures in-situ seawater fluorescence. This measurement is related to the concentration of chlorophyll in the water. But, the in-situ fluorometer must be calibrated with laboratory measurements of chlorophyll from discrete water samples. To perform this calibration we will measure the concentration of chlorophyll on some of the samples we collect with the rosette. We will first extract the pigments using acetone. Then, we will use a different type of fluorometer to measure the concentration of the extracted algal pigments. Survey methods: We will measure depth profiles of temperature, salinity, light, chlorophyll and dissolved oxygen at several locations in Bellingham Bay using a CTD. The CTD (conductivity, temperature, depth) is the oceanographer’s primary sampling device. It consists of a set of electronic probes attached to a metal rosette wheel. It will allow us to observe water properties as we lower it into the water. We will estimate salinity from conductivity. The more ions the water contains, the more electricity it will conduct. Salinity, when determined from conductivity, is usually expressed as psu (practical salinity units). The values of psu are the same as parts per thousand (%o). Temperature is expressed as degrees Celsius (oC). Other instrumentation on the CTD will measure light, dissolved oxygen, and seawater fluorescence, which is related to the concentration of chlorophyll in the water. Six water bottles can be closed to collect water at different depths as the instrument ascents using a remote electronic closing mechanism. We will collect a sample of surface water at each station and collect several water samples from several depths at the deeper stations. At shallower stations we will collect seawater using buckets, a Van Dorn water sampler, and a YSI (S, T, DO) meter. We will also collect water samples from the Nooksack River and the Post Point WWTP for nutrient analysis. Sample location: I’ve selected the sampling stations in advance based on a monitoring study I’ve been conducting in Bellingham Bay with my classes (Fig 1). Dissolved oxygen: The dissolved oxygen content of water is influenced mainly by water temperature (cold water can hold more dissolved gas than warm water) and biological activity. Primary producers (including macroalgae and phytoplankton) add oxygen to the water as they photosynthesize. Recall that photosynthesis can only take place at depths shallow enough to receive light. Aerobic organisms (plants, animals, aerobic microbes) consume oxygen during metabolism. Waters containing high levels of organic matter (i.e. dead cells, organic-rich muds, dissolved organic matter from sewage or other sources) may have low levels of oxygen because heterotrophs use up the oxygen while decomposing the organic matter. Water samples: Use a clean bucket firmly affixed to a line to obtain a surface water sample from the shallow stations. Use the Van Dorn or Niskin bottle to collect subsurface water samples (at shallow stations). Lower an open bottle to the desired depth. Send a messenger down the line to close the bottle. Pull the bottle up and remove water using the attached tubing. At deeper stations, collect water from the CTD rosette. Close (“fire’) the bottles electronically using the CTD software at selected depths. To sample the Niskin bottles, turn the knob at the top of the bottle to allow air to enter. Pull the nipple at the bottom, holding the sample bottle under the

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stream of water. Rinse the sample bottle twice with water from the Niskin bottle. Attach a glass fiber filter to the end of a 50-cc syringe. Rinse the syringe with water from the Niskin bottle and then fill it with sample water. Filter this water into the sample bottle. Also filter about 5-cc into a labeled plastic scintillation vial. Store samples in a cooler. These will be frozen for nutrient analysis during a later laboratory session. We will also collect water samples in BOD bottles to incubate for measurements of water column respiration. (This will be completed next week.) Data analysis: Generate vertical profiles of salinity, temperature, and dissolved oxygen from the stations sampled. Can you identify the thermocline, halocline, and oxycline? Calculate the density () of the seawater from the following equation of state of seawater at a pressure of one atmosphere (Gill 1982). Ignoring pressure effects makes the equation a little less accurate, but it will be sufficient for this assignment since we will be working in shallow water. A more accurate equation of state of seawater is more complicated than we can program in excel. An excel spreadsheet with the equation of state formula included is posted on the course web site.

 [kg/m3]= (999.8426 + 6.79396 * 10-2 * T - 9.0953 * 10-3 T2 + 1. 00169 * 10-4 T3 - 1.12008 * 10-6 * T4 + 6.53633 * 10-9 T5) + S * (0.82449 - 4.0889 * 10-3 T + 7.6438 * 10-5 * T2 - 8.2467 * 10-7 * T3 + 5.3875 * 10-9 * T4) + S3/2 * (-5.72466 * 10-3 + 1.0227 * 10-4 T - 1.6546 * 10-6 T2) + 4.8314 * 10-4 * S2 [T = degrees C, S = psu] Density is often expressed using the units kg/l. Convert kg/m3 into kg/l by dividing by 1000. Convert the density measurement [kg/l] into sigma-t units using the following definition: t = ((kg/l)-1) * 103 Analyzing profile data: Plot profiles of T (deg C), S (psu), dissolved oxygen, light, fluorescence and t for each station. Create a contour plot of surface salinity in Bellingham Bay. Consider the following questions: How do the profiles from the different stations compare? What might account for any differences? Which is more important in determining the density profile at the stations, temperature or salinity? How does the Nooksack River affect salinity in Bellingham Bay? Are hypoxic conditions apparent anywhere in Bellingham Bay? Where is the dissolved oxygen lowest? Draft report: Plot the locations of each station in Bellingham Bay using any program you like. Plot profiles of temperature, salinity, dissolved oxygen and density (t, calculated using formula in the provided spreadsheet. Also, create a contour plot of surface salinity. Reference: Gill, A. E. 1982. Atmosphere-Ocean Dynamics, Academic Press, San Diego, CA.

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Figure 1. Station locations for Bellingham Bay survey

Figure 2. Predicted tidal levels in Bellingham Bay, October 18, 2011.

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Figure 3. Additional chart of Bellingham Bay

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Pre-laboratory Report 3: Seawater fluorescence, chlorophyll, and phytoplankton biomass Name: _______________________________ Read the description of this week’s laboratory assignment and access the Excel spreadsheet sed_profiles.xls found on the course web site. Then, answer the following questions to be turned in at the beginning of the lab period. You do not need to type your answers to these questions. You may write your answers in the space available and turn in this sheet. 1. Calibrating an amperometric oxygen sensor: a. A bottle of seawater is in equilibrium with the atmosphere, with respect to oxygen. The seawater salinity is 31‰; the temperature is 15°C. What is the concentration of dissolved oxygen (in µM)? b. You insert the oxygen microelectrode and measure an amperage of 100 picoamperes. You measure seawater with no oxygen (bubbled with nitrogen gas). The electode reads 5 picoamperes. You insert the probe into a freshly collected seawater sample. The electrode reads 80 picoamperes. What is the concentration of dissolved oxygen in the freshly collected seawater sample? 2. A sample of seawater has a temperature of 20°C and a salinity of 32‰. What is the molecular diffusion coefficient for dissolved oxygen in this sample?

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ESCI 322 Lab Report 2 Part 2, Seawater fluorescence, light, chlorophyll, and phytoplankton biomass Primary production, timing of the spring bloom, and other oceanographic processes are affected by light intensity. Light interacts with algal pigments to drive photosynthesis; both the quality (spectrum, or color) and quantity of light are important in regulating this process (Fig. 1). The objectives of this laboratory session are to try several different methods for the measurement of irradiance (quantity of light). We will examine the interaction of light with algal pigments, including properties of light absorbance and fluorescence, and we will use pigment fluorescence to estimate concentrations of algal chlorophyll in samples collected from Bellingham Bay.

Fig. 1 Electromagnetic spectrum. The area under the lower curve represents the total energy received from the sun, divided into the proportions received in the form of UV, visible (colored) and infrared wavelengths. From Thurman (1997).

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1) Methods for measuring irradiance and seawater fluorescence Secchi disk: A black and white disk is a low-tech way to measure the penetration of light into water. Named for Father Pietro Angelo Secchi, this primitive instrument has been used extensively in marine and aquatic systems for studying irradiance. Today, when the quantum sensor smashes on the side of the ship, the Secchi disk comes out. Lower the disk until it can no longer be seen, then raise it to the depth at which it first becomes visible. Record this depth. You may have to repeat this several times to obtain a consistent Secchi depth. Spherical light sensor (4pi): This spherical light sensor measures the total amount of photosynthetically active radiation (PAR, radiation at wavelengths that can be used in photosynthesis). PAR wavelengths range from 400 to 700 nm for most photosynthetic organisms. Although light enters the water from above, it is scattered by water molecules so that, from an underwater vantage point, light comes from all directions. The spherical (4pi) sensor allows accurate measurement of this diffuse light field. Irradiance is reported in units of µmoles photons m-2s-1. Obtain an irradiance profile by lowering the sensor (carefully!) to discrete depths as indicated by markings on the cable. (Note that the same data logger can be used with a 2 pi [flat, circular] sensor to measure the amount of light striking the water’s surface, either instantaneously, or integrated over time.) In situ fluorometer: Chlorophylls absorb light energy at one wavelength and emit it a longer wavelength; this property is known as fluorescence. The fluorometer works by shining blue light onto a sample (called excitation) and measuring the resulting emission of red light. Filters are used to control the wavelengths received by the sample and the detector (a photomultiplier tube). An in situ fluorometer measures the fluorescence of whole seawater. By picking the appropriate excitation and emission wavelengths, the measured seawater fluorescence should be related to the amount of chlorophyll in the water. However, other water properties also fluoresce. Therefore, the fluorescence data must be calibrated by comparing them to measured chlorophyll concentration in discrete samples. 2) Collection of water samples for chlorophyll and (later) nutrient analysis. Determine the depth of the pycnocline and the depth of maximum fluorescence using data collected electronically by the CTD. Then, collect water samples from six depths using the water sampling bottles on the CTD. Collect samples from above and below the pycnocline and at depths with different fluorescence levels. Once on deck, remove water from the CTD rosette bottles and fill one plastic sample bottle with water from each bottle; label with depth and station ID. Filter the samples on deck, store in aluminum foil, and place into a cooler. These samples will be frozen for later analysis in the laboratory. 3) Calculation of extinction coefficients From Secchi disk data: Light in water is absorbed and scattered by the water molecules themselves, and by dissolved and particulate material in the water. The amount of light absorbed per unit surface light per

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meter depth is given by the extinction (or attenuation) coefficient k (m-1). The light attenuation coefficient, KD, is calculated from the Secchi depth (ZSD) according to the formula KD=1.44 /ZSD. From light meter data: The extinction coefficient KD can also be calculated from vertical profiles of irradiance such as those you obtained using the light meter. In this case, light (I) is assumed to decrease exponentially with depth (z) according to the formula IZ = I0 e-kz. Based on this model, the extinction coefficient can be calculated from a near-surface (I0) and a deep (IZ) irradiance measurement and the depth interval (z) between these two measurements: k = -1/z ln(IZ/I0). Note that the shallowest irradiance measurements are often ‘noisy’, so you probably will not want to use these for this two-point method of calculating k. The most accurate way to estimate k is to use all your data from a given vertical profile. Create a plot of ln irradiance vs. depth (analogous to a semi-log plot for estimating growth rate). The slope of the regression of ln irradiance vs. depth is -k. 4) Light absorption by algal pigments All photosynthetic organisms contain pigments (Fig. 2) to harvest light energy and to protect themselves from light-induced damage. Spectrophotometry takes advantage of light absorption by pigments to estimate their concentration in a given sample. Within a certain range of concentration, the absorbance of light is proportional to the concentration of pigment present (Beer’s Law). The spectrophotometer passes a beam of light through a substance (in our case, an organic extract) and the amount of light absorbed from the beam by the sample is determined. The photo-diode array spectrophotometer is able to quantify absorbance over a range of wavelengths simultaneously. We will examine absorption spectra from several different kinds of pigments using the diode array spectrophotometer, including extracts of the pigments from our study site. This kind of spectrophotometry can used quantitatively to determine pigment concentrations in a water sample (see Parsons et al 1984 and Jeffrey et al. 1997 for methodologies).

Fig. 2. Diagram of a cryptophyte cell. These are common members of marine phytoplankton communities. The large chloroplast is bounded by membranes (CE, CER) and filled with layered thylakoid membranes (T). Most of the pigments are imbedded in the thylakoid membranes (Lee 1999).

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Fig. 3. Absorbance spectra of some commonly occurring chlorophylls and carotenoids. 5) Measurement of pigment concentrations by fluorometry Chlorophyll a fluorescence can be used to quantify the amount of chlorophyll present in the particles in a water sample. This method is very sensitive, so it works well for dilute systems such as the ocean. The fluorometer works by shining blue light onto a pigment extract and measuring the resulting emission of red light. Filters are used to control the wavelengths received by the sample and the detector (a photomultiplier tube). The amount of blue light used to excite the fluorescence will influence the amount of fluorescence produced; this is controlled by a series of “doors” and must be accounted for in the calculations. The fluorometer is standardized using pure chlorophyll a extracts which in turn are quantified on the spectrophotometer (this has been done for you). There are four steps involved in the measurement of water column chlorophyll concentrations: i) filtering the water sample; ii) grinding the filter (and attached particles) in acetone; iii) measuring the fluorescence of the sample in a fluorometer; iv) calculating chlorophyll concentrations from fluorescence readings. Step 1: Filtering the water sample a) Use labeling tape to label a set of 15-ml centrifuge tubes with your sample names. b) Place a clean glass fiber filter in a plastic threaded filter holder; close the filter holder and attach it to a 50-ml syringe with plunger removed. c) Gently invert your water sample several times to resuspend and mix the particles. Measure 50 ml of the sample in a graduated cylinder; pour this into the syringe and gently but firmly push the water through the filter with the plunger. Catch the filtered water in a small plastic sample bottle

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(this will be saved for nutrient analysis). d) Label the nutrient sample bottle with the sample name. e) Place the filter in the appropriate 15-ml centrifuge tube. This filter now has on it all the particles (including chlorophyll-containing phytoplankton cells) that were in the original 50-ml water sample. Step 2: Sonicating the filter a) Place a filter from step i into a 15-ml centrifuge tube and add 5 ml cold 90% acetone. b) Sonicate while submerging the centrifuge tube in an ice-water bath for one minute. Wear gloves, safety goggles and ear protection. c) When the sample is thoroughly sonicated, add acetone until the final volume is 10 ml. d) Record the final volume of solvent plus homogenate in the tube. This is your “extraction volume”. Put the tube into a test tube rack for storage in the ice bath (or freezer for longer-term storage). Step 3: Measuring the sample fluorescence a) Vortex or vigorously shake each centrifuge tube, then remove the filter, squeezing out any solvent using a clean (solvent-rinsed) pair of forceps. Centrifuge the tubes (high speed, 5 min). b) If extracts are visibly green, they must be diluted or the detector response will be saturated. Use calibrated centrifuge tubes and automatic pipettes to dilute samples with 90% acetone; keep track of all dilutions. c) Zero the fluorometer using a cuvette containing 90% acetone. Re-zero every time you switch door (sensitivity) settings. d) Transfer your extract to a clean glass cuvette. Be careful not to resuspend any of the palletized filter debris (this will interfere with the fluorescence reading). e) Read the fluorescence of your extract. This value should be >25 and H4SiMo12O40 + 12 H2O (Silicomolybdic acid)

Wait at least 15 minutes before proceeding to the next step. This reaction forms a yellow compound (silicomolybdic acid), whose concentration is equal to that of the silicic acid initially present in the sample (1:1 stoichiometry). In samples with high [Si(OH)4] you will be able to see a pale yellow color form.

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6. Mix reducing reagent: For each 2.5-ml sample to be analyzed mix 0.2 ml Metol-sulfite reagent, 0.12 ml 10% oxalic acid, 0.12 m1 50% sulfuric acid and 0.16 ml DIW for a total volume of 3.1 ml. Use polypropylene graduated cylinders. First, rinse each cylinder twice with a few ml of Metol-sulfite reagent. Multiply these volumes by the number of samples to determine the total volume required for each reagent (see table below). Measure the reagents into a plastic beaker and cover with parafilm. Reagent Metol-sulfite Oxalic Acid H2SO4 DIW

1 sample 0.2 ml 0.12 ml 0.12 ml 0.16 ml

10 samples 2 ml 1.2 ml 1.2 ml 1.6 ml

20 samples 4 ml 2.4 ml 2.4 ml 3.2 ml

50 samples 10 ml 6 ml 6 ml 8 ml

100 samples 20 ml 12 ml 12 ml 16 ml

7. Add 1.5 ml of the mixed reducing reagent to all samples and standards. Cap tightly and shake once or-twice. Wait at least 2.5 hr for the yellow silicomolybdic acid to be reduced to a deep blue "silicomolybdous acid complex" - whose exact formula isn't known. In samples whose [Si(OH)4] is < ~1.5 M it will be difficult to see any color, but all others should be visibly blue. (If samples are not visibly blue, the waiting period can be reduced to 1.5 hr.) While you are waiting for this reaction to occur, proceed to the phosphate measurements. 8. Now, add 1.0 ml of acidified ammonium molybdate reagent to the blank. This procedure is called a reverse-addition blank. The oxalic acid in the mixed reducing agent blocks the formation of the yellow silicomolybdic acid so none forms when the acidified ammonium molybdate reagent is added later. 9. Measure the absorbance at a wavelength of 810 nm in a spectrophotometer (the blue complex absorbs most strongly in the infrared), using glass culture tubes. We can use glass at this point because the silicate that we will measure has already formed the blue silicomolybdous acid complex and the oxalic acid blocks further complex formation. Calculations: The [Si(OH)4] of each sample is calculated as: [Si(OH)4] (M)

= Sample absorbance - Reverse-addition blank absorbance Slope of standard curve (absorbance/M)

I will provide you with an excel spreadsheet that will help you make the appropriate calculations.

Dissolved phosphate (H2PO4-, HPO42-, PO43-) determination (Parsons et al. 1984) Phosphate is an important limiting nutrient in freshwater and marine systems. Eutrophication in freshwater systems are often the result of high inputs of P, which lead to increased growth of algae, and sometimes to reductions in dissolved oxygen or other problems. The importance of phosphate for regulating the growth of algae was spectacularly demonstrated by Vollenweider (1976). Figure 1 shows the relationship between total P loading and algal biomass in lakes.

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Figure1. Empirical model relating mean surface chlorophyll-a concentrations of lakes to phosphorus loading, adjusted for hydraulic loading. Plot from Cloern, 2001. Data from Vollenweider, 1976.

Outline of method: The seawater sample is allowed to react with a reagent containing molybdic acid, ascorbic acid, and trivalent antimony. The resulting complex is reduced to give a blue solution that is measured at 885 nm. Procedures: Samples should be at room temperature. Turn on Spectrophotometer and set wavelength to 885 nm. This is the light absorption maximum for the blue solution produced in this method. Standards: To save time we will make a set of working standards from a secondary standard that has been prepared in advance. First a primary standard solution was made from anhydrous KH2PO4 and DIW. The secondary standard was made from the primary standard. Label and rinse five 100-ml volumetric flasks several times with DIW. Fill them with about 90 ml of DIW. Then, rinse an Erlenmeyer flask several times with a few ml of the secondary standard solution. Using a 5-ml pipette, carefully add the secondary standard solution from the Erlenmeyer flask to each 100-ml volumetric flask according to the table below. Bring up the volume to 100 ml by carefully adding DIW. These are your tertiary standards. 3° STD (µM) 0.0 0.6 1.8 3.0 4.2

ml 2° STD / 100 ml H2O 0.00 0.50 1.50 2.50 3.50

Place ten glass culture tubes in a tube rack (two for each standard) and record their positions in your notebook. Also add five tubes for the seawater sub samples and record their positions. ECSI 322 – Oceanography Laboratory - Manual

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Mixed reagent: We will create the mixed reagent from single reagents that were made in advance of today’s lab. For each 5-ml sample, use 2 ml Ammonium molybdate, 5 ml Sulfuric acid, 2 ml Ascorbic acid, and 1 ml K-antimonyl-tartrate. First, wash a graduated cylinder with a few ml of ammonium molybdate three times. Then, measure the correct volume of ammonium molybdate and the other reagents according to the table below. Add these reagents to a labeled Erlenmeyer flask, cover with parafilm, and mix for 30 seconds. Use volumes in the 50 samples column. Reagent Ammonium molybdate Sulfuric acid Ascorbic acid K-antimonyl tartrate

10 samples 2.0 ml 5.0 ml 2.0 ml 1.0 ml

20 samples 4.0 ml 10.0 ml 4.0 ml 2.0 ml

50 samples 10.0 ml 25.0 ml 10.0 ml 5.0 ml

Measure 5 ml of each sample, each STD, and the nanopure H2O blank into culture tubes. Add 0.5 ml mixed reagent. Wait 5 minutes. Measure absorbance on a spectrophotometer. Spectrophotometric Procedures Set the spectrophotometer at 885 nm. Turn spec on and allow to warm up for 15 min. Set wavelength to 885 nm. Set filter lever to the right. Set to 0% transmittance. Change mode to “absorbance”. Insert Nanopure water blank (this is not the same as the reagent blank). Set to 0% absorbance. Insert sample and record absorbance. Data Analysis for Phosphate Follow the data analysis procedures for silicate. Reporting nutrient concentrations in Bellingham Bay The spatial distribution of temperature, salinity, and nutrients changes every time I sample Bellingham Bay with students from this class. This is because each time we sample it at different stages of the tide and under different wind conditions, and because the Nooksack River flow varies from year to year. One way to make sense of the nutrient concentrations we measure, however, is to plot them versus salinity. This enables us to examine the relationships between river input from the Nooksack, salt water input from the Strait of Georgia and the Strait of Juan de Fuca, and nutrient concentrations without reference to specific locations within the bay. We can even infer whether biological or chemical processes produce or consume nutrients within Bellingham Bay as well. This method will be described next.

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Understanding relationships between salinity and nutrient concentrations

Tracer concentration

A

River source

Tracer concentration

Mixing theory: A non-reactive solute (sometimes referred to as a conservative tracer) input from a river will be diluted by seawater. If dilution is the only process that determines the concentration on such a tracer in an estuary, the relationship between tracer concentration and salinity will be linear. This also holds for a tracer with a seawater input.

Salinity

B Marine source

Figure 1. Theoretical relationship between a conservative solute and salinity for solutes with (A) a river source and (B) a marine source.

Salinity

Tracer concentration

A non-conservative tracer is one that has a source or sink within the estuary. In this case, the relationship between tracer A B concentration and salinity will Estuarine source be non-linear.

Salinity

Salinity

Estuarine sink

Figure 2. Theoretical relationships between nonconservative solutes and salinity for tracers with a river source (A) and a marine source (B).

By plotting nutrient concentrations versus salinity, we can infer whether the major sources or nutrients to Bellingham Bay are from rivers or deeper, high salinity marine waters entering the bay at depth. Deviations from a linear relationship indicate sources or sinks of N and P in the Bay. Like salinity, the Redfield ratio can also be used to infer something about nutrient sources and sinks in Bellingham Bay. If the DIN:DIP ratio deviates greatly from ~16, that would suggest a source or sink of N or P. Although there are many processes that could cause nutrient concentrations to deviate from Redfield proportions, the variation in this ratio with salinity can be used to learn something about nutrient cycling processes.

DIN:DIP ratio

N > P: Nitrogen fixation?

16

8

N > P: Phosphate storage In sediments?

Redfield ratio

N < P: Fertilizer input?

N < P: Denitrification In sediments?

Figure 3. Several processes that can cause nutrient concentrations to deviate from Redfield proportions. (This list is not complete.) Deviations at low salinity may reflect river inputs. For example, phosphate-based fertilizers reduce the N:P ratio whereas freshwater N fixation can increase it. Deviations at high salinity in an estuary may reflect processes affecting bottom waters. For example,

0 Salinity

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denitrification in marine sediments can reduce N:P whereas sorption of P onto sediments could potentially increase the ratio. Also, preferential uptake of N versus P in the marine waters entering Bellingham Bay could reduce the N:P ratio. Creating a nutrient budget for Bellingham Bay: The data we have collected will allow us to calculate a nutrient budget. This is a careful accounting of the rates of supply of nutrients from different sources. The four potentially important sources that we can compare are: the Nooksack River, the Post Point WWTP, nutrient supply from sediment, and deep water inflow from the Strait of Georgia/Strait of Juan de Fuca. For the sediment component of the budget, divide the nutrient flux by the water depth as we did for the sediment oxygen consumption rate. For the other components, we need to multiply the nutrient concentrations in water from the Nooksack River, Post Point WWTP, and deep water with their volumetric flow rates. Example: (15 mmole/m3 Nitrogen) * (7,000,000 m3/day Nooksack R. water) = 1.05*108 mmole/day nitrogen from the Nooksack River. Create a budget for both nitrogen and phosphorus. You’ll need to calculate total nitrogen as the sum of ammonium, nitrate and nitrite. Flow rates for the Nooksack River are available from the USGS. Flow rates for the Post Point WWTP are available too. We can calculate the volumetric flow rate of deep water from the Nooksack River flow rate and the extent to which Bellingham bay water is diluted. Formula: Deep-water inflow = R[SO/(Si-SO)], where R is the Nooksack River flow, Si is the salinity of incoming deepwater and SO is the salinity of outgoing surface water. Report: Our data on nutrient concentrations and salinity from various locations throughout Bellingham Bay will allow us to ask the question: What are the sources and sinks of nutrients in Bellingham Bay? To address this overall question, consider the following auxiliary questions. What is the relationship between salinity and nutrient concentrations in Bellingham Bay? Is the Nooksack River an important source of nutrients to Bellingham Bay or are higher salinity bottom waters more important? According to your nutrient budget, which source is the most important? Are there other potential sources of nutrients that we have overlooked? Are nutrients conservative tracers or do they have sources or sinks in Bellingham Bay? Do nutrient concentrations in Bellingham Bay follow Redfield ratios? If not, what does the deviation from Redfield proportions tell you about nutrient cycling in Bellingham Bay and Puget Sound? Address these questions by plotting your data following Figures 1-3 and compare your data with theoretical expectations. Present your nutrient budget as a table in your report. References Cloern, J. E. 2004. Our evolving conceptual model of the coastal eutrophication problem. Mar. Ecol. Prog. Ser. 210, 223-253. Parsons, T. R., Y. Maita, and C. M. Lalli. 1984. A manual of chemical and biological methods for seawater analysis. Pergamon Press, Elmsford, N.Y. Strickland, J. D. H. and T. R. Parsons. 1972. A Practical Handbook of Seawater Analysis. Fish. Res. Bd. Can. bull 167 . 2nd ed. pp 65-70. Vollenweider, R. A. 1976. Advances in defining critical loading levels of phosphorus in lake eutrophication. Mem. Ist. Ital. Idrobiol. 33, 53-83.

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Pre-laboratory Report 5, Phytoplankton growth and grazing Name: _______________________________ Read the description of this week’s laboratory assignment and answer the following questions to be turned in at the beginning of the lab period. You do not need to type your answers to these questions. You may write your answers in the space available and turn in this sheet. But, you’ll also need to turn in a data plot and linear regression described below. Problem: You conduct a dilution experiment. After making the dilution series shown below, you allow phytoplankton to grow for 24h and then stop the experiment by filtering the phytoplankton. You measure the concentration of chlorophyll-a in the filtered samples. The data you collect are shown in the table below. Table. Hypothetical results from a dilution experiment Dilution factor (fraction seawater) 1.00 0.75 0.50 0.25 0.05

Initial chlorophyll concentration (p0) 30 22 11 3.0 0.15

Final chlorophyll concentration (p) 38.5 53.3 47.1 29 4.0

Net growth 1/t * ln(p/p0)

1. Fill in the last column of the table [ 1/t ln(p/p0) ]. 2. Calculate the slope and intercept from the linear regression of the net growth versus dilution factor. (Show the Excel plot you created including the data points and linear regression line.) Slope: _____________ Intercept: __________ 3. What are the per-capita growth and grazing rates that you determined from this experiment? Growth rate: ________________ Grazing rate: ________________

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ESCI 322 Lab Report 4, weeks 9 and 10: Phytoplankton population growth and grazing Part 1: Setting up the experiment The abundance of phytoplankton in the ocean is set by the balance between rates of population growth, mortality and transport. Phytoplankton population growth rates in the field are affected by light, nutrient concentration, temperature, and species composition among other factors. Consumption by grazers is one of the largest sources of mortality. There are many grazers of phytoplankton in Puget Sound; herring, calanoid copepods and bottom-dwelling filter feeders such as clams are relatively large grazers. There are also phytoplankton grazers that are barely larger than the phytoplankton cells they consume. These protistan grazers are termed microzooplankton. In many marine ecosystems, microzooplankton are extremely important phytoplankton consumers. The objective of this lab is to measure rates of phytoplankton growth and microzooplankton grazing. We’ll perform the experiment at two light levels to better understand how light and grazing interact to control phytoplankton abundance in the sea. Theory: Consider the equation for population growth. It can be written in differential form as dp follows:  p , where p is the concentration of phytoplankton and µ is the per-capita growth dt rate. If z is the microzooplankton concentration and χ is the phytoplankton-zooplankton dp encounter rate, you can add grazing to this equation as follows:  p  zp . Now, consider dt what would happen if you diluted a sample of seawater with filtered seawater. Dilution should not affect the growth rate of phytoplankton which multiply by binary fission. However, dilution will reduce the encounter rate between phytoplankton and zooplankton, lowering the grazing rate. If you add dilution (D is the fraction of undiluted seawater in the sample.) to the population dp growth model, you can write it as  p  gDp , where g is the per capita mortality rate of dt phytoplankton due to grazing. The solution to this model is p = p0e(µ-gD)t, where p0 is the initial phytoplankton concentration. Taking the natural log of both sides and rearranging this equation 1  p      gD . The first term is a measure of phytoplankton gives the following formula: ln  t  p 0  net growth rate. Plotting this versus the dilution factor, D, yields a straight line with a slope equal to g and intercept µ (Fig. 1). Thus, if you conduct a dilution experiment you can calculate both the phytoplankton per capita growth and grazing rates. In our experiment, we will conduct a dilution experiment with two light intensity treatments (full-strength light and 50% light). We’ll use mesh screening material to manipulate light intensity and measure phytoplankton growth and grazing rates at the two light levels. This will allow us to ask the question how does phytoplankton growth respond to light and grazing? Methods: It is a challenge to work with phytoplankton in late fall when biomass approaches its lowest levels due to low light levels. So, I will collect a big sample of seawater and let the algae grow under well-lit conditions in the laboratory for a week prior to our class. That should give us a sample with a high enough biomass to use for our experiment. Filter the water to create three dilution levels – full-strength seawater, 50% seawater diluted with filtered seawater, and ECSI 322 – Oceanography Laboratory - Manual

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10% seawater diluted with filtered seawater. Perhaps the trickiest part of setting up this experiment will be filtering the water. You’ll need to filter a large quantity of water without damaging the phytoplankton cells and then divide the seawater into replicate bottles while keeping the cells in suspension so that all bottles receive the same concentration of cells. Filter half the seawater through a filter cartridge using a peristaltic pump which won’t damage the plankton. While filtering, use a piston to keep the cells in suspension. Then, combine the whole and filtered seawater to create the following treatments, keeping the cells in suspension while doing so. Fraction whole Seawater 100% 100% 100% 50% 50% 10% 10% 10%

Number of replicates 4 2 2 3 1 4 2 2

Light level NA 100% 10% NA 100% NA 100% 10%

Sampling time initial final final initial final initial final final

Incubate the samples for 48 hours. End the experiment by preserving a 15-ml sample from each bottle using Lugol’s iodine preservative and pass the rest of the samples through a glass fiber filter. Freeze the filters for analysis next week.

Dilution experiments Y-intercept= “infinite dilution”

Net Growth Rate (per time)

GROWTH RATE

Growth equation: dp/dt = p – zp dp/dt = p – zDp = p – gDp Solution: 1/t ln(p(t)/p0) =  – gD

Slope = linear relationship with dilutions HERBIVORY RATE

Decreasing # of Grazers Dilution factor (fraction SW) 0

1

Figure 1. Theoretical relationship between net growth rate and dilution factor. The slope gives the per capita grazing rate and the y intercept gives the per capita growth rate. Reference: Landry, M.R., 2001. Microbial loops. In: Steele, J.H., Thorpe, S., Turekian, K. (Eds.), Encyclopedia of Ocean Sciences, Academic Press, London, pp. 1763–1770.

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ESCI 322 Lab Report 4, weeks 9 and 10: Phytoplankton population growth and grazing Week 2: Sample processing and analysis Reconsider the population growth equation that forms the basis of our dilution experiment: 1  p      gD . Note that ln(p/p0) is unitless. It depends on the ratio of p/p0 (the ratio of ln  t  p 0  final to initial phytoplankton concentration) and the units cancel. This means we can use any measure of concentration we like to quantify the phytoplankton. The units don’t matter. We will use chlorophyll concentration since it can be measured relatively easily and precisely. We will also enumerate phytoplankton and microzooplankton cells to determine which species were most strongly affected by grazers. Measuring chlorophyll concentration by fluorometry: Chlorophylls absorb light energy at one wavelength and emit it a longer wavelength; this property is known as fluorescence. Fluorescence measurements are quite sensitive so it can be used to measure chlorophyll in dilute systems like ours. The fluorometer works by shining blue light onto a pigment extract and measuring the resulting emission of red light. Filters are used to control the wavelengths received by the sample and the detector (a photomultiplier tube). The amount of blue light used to excite the fluorescence will influence the amount of fluorescence produced; this is controlled by a series of “doors” and must be accounted for in the calculations. The fluorometer is standardized using pure chlorophyll a extracts which in turn are quantified on the spectrophotometer (this has been done for you). There are four steps involved in the measurement of water column chlorophyll concentrations: i) filtering the water sample; ii) grinding the filter (and attached particles) in acetone; iii) measuring the fluorescence of the sample in a fluorometer; iv) calculating chlorophyll concentrations from fluorescence readings. Step 1: Filtering the water sample I have already filtered the samples and have frozen them for today’s analysis Step 2: Sonicating the filter a) Place a filter from step 1 into a 15-ml centrifuge tube and add 5 ml cold 90% acetone. b) Sonicate while submerging the centrifuge tube in an ice-water bath for one minute. Wear gloves, safety goggles and ear protection. c) When the sample is thoroughly sonicated, add acetone until the final volume is 10 ml. d) Record the final volume of solvent plus homogenate in the tube. This is your “extraction volume”. Put the tube into a test tube rack for storage in the ice bath (or freezer for longer-term storage). Step 3: Measuring the sample fluorescence a) Vortex or vigorously shake each centrifuge tube, then remove the filter, squeezing out any solvent using a clean (solvent-rinsed) pair of forceps. Centrifuge the tubes (high speed, 5 min). b) If extracts are visibly green, they must be diluted or the detector response will be saturated. Use calibrated centrifuge tubes and automatic pipettes to dilute samples with 90% acetone; keep

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track of all dilutions. c) Zero the fluorometer using a cuvette containing 90% acetone. Re-zero every time you switch door (sensitivity) settings. d) Transfer your extract to a clean glass cuvette. Be careful not to resuspend any of the palletized filter debris (this will interfere with the fluorescence reading). e) Read the fluorescence of your extract. This value should be >25 and