envr 322 oceanography laboratory

ECSI 322 – Oceanography Laboratory - Manual 1

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 17, 2014

mailto:[email protected]


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 smaller lab

assignment, and “pre-laboratory” assignments. Each assignment will be typed, although figures

and graphs may be drawn by hand. The reports 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 will be handed in first for evaluation and comments as a component of

pre-laboratory reports. Comments on the draft of the results section are intended to aid students

in the completion of the final reports. 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 first assignment 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 80%

Pre-lab assignments 20%

Approximate grading scale:

93-100 A 90-92 A- 88-89 B+ 83-87 B 80-82 B- 78-79 C+

73-77 C 69-72C- 67-68 D+ 61-66 D 57-60 D- 0-56 F


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 will not be accepted late.

LAB REPORT FORMAT

A laboratory report is a document in the form of a scientific paper. Writing 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 science 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, 2014.

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.

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, methods, 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.

One way to organize an abstract is to simply write a sentence or two summarizing your

introduction (why you did it), methods (what you did), results (what you observed), and

discussion (what it means).

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). You must cite published papers from the peer-reviewed

literature in your introduction. Internet sites are rarely cited in the introduction.


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 an exhaustive literature

review. Keep the introduction focused on the problem you are addressing.

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.

The methods section is NOT a sequential recounting of everything you did. It does not tell

a story or go into the nitty-gritty details unless these are important for the reader to know

in order to repeat the work. Instead, it works better if you organize your methods sections

around specific tasks. For example, "Concentrations of nutrients (ammonium, nitrate+nitrite,

and phosphate) were measured following the methods of Parsons et al. (1984). Samples were

collected from Niskin bottles into clean and sample-water rinsed 100-ml polyethylene bottles.

Sample water was first filtered through a gf/c glass-fiber filter (nominal diameter 0.7-μm) and

was frozen prior to analysis. Ammonium was measured on thawed samples as follows..." The

idea is to briefly tell the reader the important details of your methodology. The following

structure is NOT acceptable: “First we collected water samples from a Niskin bottle. We then

filtered the samples through a gf/c glass fiber filter. Samples were frozen for one week and then

the following week we measured nutrient concentrations in the analytical lab at the Shannon

Point Marine Center.” The problem with the second example is that it includes information that

is not critical such as the location where samples were analyzed. (It is not necessary for all

marine scientists to measure their nutrient samples at SPMC.) It also describes the activities

chronologically as though the order in which the activities were performed was critical.

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.

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.) Results sections are much more fun to read if you use this

section to tell the reader a story about what you found. Although most of the text will describe

the results summarized in figures and tables, try to be interesting as you lead the reader through

your discoveries.

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 y-

in axis. (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 standard deviation) 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 the 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 readers

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. This means you must cite published papers in your

discussion. You may also want or need to discuss short-comings in 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. Example 1: Several other species of

anemone are known to have respiration rates that increase with temperature (Matthews, 2003;

Smith and Wesson, 2014). Example 2: Our findings of lower respiration rate at lower

temperatures agreed with those of Matthews (2003). 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.


ESCI 322 Lab Assignment : Waves and Coastal Processes

Objectives: 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.

Broader learning objectives: Compare measurements with theory and evaluate their agreement.

Develop hypotheses as a group and then conduct a laboratory experiment to test them. Begin to

develop skill using Microsoft Excel as a tool for analyzing and displaying data.

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.

2

1

2tanh

2

L

dgLC

,

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 = 2gL (m/s). Since g is a

constant, this formula reduces to C= L25.1 (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 = d13.3 (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.

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 – In order to gather the information necessary for your report, pay close attention to

the underlined questions and activities during the lab session.

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?

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: 2

1

2tanh

2

L

dgLC

Properties of breaking waves: 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?

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 Wavelength


Assignment summary

Rather than writing a formal report, do the following. This assignment will be due in one week.

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?


Pre-laboratory Report 3: Coastal sediments and benthic communities of

Burrows Bay

Name: _______________________________

Read the description of this week’s laboratory assignment. 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.

Sediment grain size:

Sediment grain diameters are classified according to the logarithmic phi scale. Calculate the phi

sizes of the following:

Sediment grain with a diameter of 0.1 mm: __________________ phi

Bacterium with a diameter of 1 mm: _________________ phi

The earth with a diameter of 12,720 km : ________________ phi

Remember the following when you answer the above questions:

1 μm = 10-6 m

1 mm = 10-3 m

1 km = 103 m


Lab Report 4: Coastal sediments and benthic communities of Burrows Bay

The objective this lab is to collect sediment samples from Burrows Bay and to observe changes

in sediment properties and faunal communities along a transect from Alexander Beach toward

the Allan Pass.

Sampling Methods

We will collect our samples using a Peterson grab. This is perhaps the simplest of all ship-

deployed benthic samplers. We will reserve some of the sediment for grain-size analysis and we

will wash the rest of the material through a 1-mm mesh sieve and look for animals. At each site

we will note the water depth and determine location by GPS.

Questions to consider during our sampling trip

1: How do sediment properties vary along our transect? What is the source or coarser-grained

sediments? What is the source for the finer sediments? How patchy is it? What could

account for these patterns?

2: What kinds of animals do we observe at each sampling site? Can you identify any patterns of

faunal community change related to sediment grain size?

Sample processing

Animals

The faunal samples will be fixed in formalin for 5 d, transferred to ethanol and stained with Rose

Bengal. We will later pick out the animals under dissecting microscopes and identify them to the

lowest possible taxon. We will then divide the organisms into the following trophic groups:

suspension feeders, deposit feeders, and predators.

Sediment grain size

Sediments can be classified according to size by the Wentworth scale.


Note that the Wentworth scale groups sediments into logarithmic size classes. The unit used to

classify sediment size on a log scale is called phi. Phi = -log2(diameter in mm). We will

separate sediments into the following size classes:

Sediment Type Size in mm Phi = -log2(diam. [mm])

Gravel > 2 -1

Sand Very coarse >1 0

Coarse > 0.5 1

Medium > 0.25 2

Fine > 0.125 3

Very fine > 0.0625 4

Silt > 0.032 5

clay > 0.016

> 0.008

< 0.008

6

7

8

The grain-size samples will be processed the week after the sampling trip. First, we will add

~100 ml of distilled water to each sample jar and shake them. We will then pour the sample

through a 63-μm sieve set inside a large funnel, and collect the water in a 1000-ml graduated

cylinder. Rinse the sediment in the sieve and funnel thoroughly with a squirt bottle and fill the

graduated cylinder with water. Cap with Parafilm. Transfer the sediment retained by the 63-μm

sieve onto a piece of labeled filter paper in vacuum apparatus. Vacuum for five minutes. Place

sample in drying oven overnight. After drying, we will separate the > 63-μm fraction into grain-

size classes. The < 63-μm fraction will be added to the graduated cylinder and smaller size

fractions will be separated by gravitational settling velocity. Top off the graduated cylinder to

make the volume 1000 ml.

To separate the finer sediment into size fractions, overturn the graduated cylinder (keeping a

tight grip on the end covered in parafilm) for three minutes. Then, set the cylinder down and

immediately collect a 20-ml sample from a depth of 20 cm. Dispense the sample into a pre-

weighed glass beaker. Take similar 20-ml samples at the following intervals: 5 min 32 sec, 22

min 4 sec, and after 1 hr 28 min. Dry the beakers in the drying oven, then weigh the beakers and

subtract the beaker mass to calculate the mass of dry sediment. Fill in the table and calculate the

mass of sediment in each size fraction.

Preview of report (to include data from this week’s lab and next week’s)

The lab report will be formatted as a scientific paper (see syllabus for detailed description of the

format). The introduction to the report should discuss the relationship between animals and

sediments. See Sanders (1958) for a an old but classic paper on this topic. The methods and

materials section should include a chart of Burrows Bay with our sampling stations plotted on

the chart. You can use Google maps to generate your plot, but beware that the satellite view is a

little too busy for a report figure. It should also describe the way the grain-size and faunal

samples were collected and processed.


Results section

Calculate the median grain size for each sampling site. Display the size distribution of each

sample by creating a column chart. Plot the sediment mass for each size fraction as columns and

use the phi classes as categories. E.g.,

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

-1 0 1 2 3 4 5 6 7 8

phi

fracti

on

(b

y m

ass)

Display the proportion of animals from different trophic groups at each site. Choose an

appropriate graph for this (pie chart, column chart, or other chart of your choice).

References

Rhoads, D.C. and Young, D.K. 1970. The influence of deposit-feeding organisms on sediment

stability and community trophic structure. J. Mar. Res. 28: 150-178.

Sanders, H.L. 1958. Benthic studies in Buzzards Bay. I. Animal-sediment relationships.

Limnology and Oceanography 3: 245 – 258.

Snelgrove, P.V.R. and C.A. Butman. 1994. Animal-sediment relationships revisited: Cause

versus effect Oceanogr. Mar. Biol. Ann. Rev. 32: 111-177.


Pre-laboratory Report 4: Coastal sediments and benthic communities of

Burrows Bay

1. Create a chart of Burrows Bay and plot our station locations on the chart.

2. Read the paper by Sanders (1958) and answer the following:

The proportion of deposit-feeding and suspension-feeding benthic organisms varies with

sediment grain size, which contributes to spatial variation in benthic communities. List two

processes that might account for the variation in benthic communities with sediment.


Lab report 4 continued: Coastal sediments and benthic communities of

Burrows Bay II – Macrofaunal feeding modes

The objective of this lab is to identify organisms from the samples we collected along the

transect from Alexander Beach to Allan Pass.

Sample processing

Animals

The faunal samples were fixed in formalin for 5 d and were then transferred to ethanol and

stained with Rose Bengal. Today will later pick out the animals under dissecting microscopes

and attempt to identify them to the level of family. We will then divide the organisms into the

following trophic groups: suspension feeders, deposit feeders, and predators. Most of the

organisms we will find will be polychaetes and bivalves. We will also likely find gastropods,

amphipods, and other crustaceans. The feeding mode of the polychaetes can generally be

determined by identifying the organism to the level of family. For the bivalves, the deposit

feeders will all belong to the order Nuculoida or to the family Tellinidae. All others will be

suspension feeders.

Sediments

We will also finish weighing the sediments from the various size fractions and use the excel

spreadsheet to calculate the proportion of sediment in each size class. Also calculate the median

grain size and the percent silt-clay for each sample.

Report

Details of the report format were given in Week 1, Coastal sediments and benthic communities

of Burrows Bay. Again, the report should address the following questions:

- Why does sediment grain size vary along our transect?

- How does the proportion of animals in different trophic groups vary among our

stations?

- What processes might cause this variation?

In the discussion section, you should describe a hypothesis to explain the patterns we observed

and suggest a way to test this hypothesis.

The following references will help you as you think about the relationships between organisms

and sediment grain size.

References

Rhoads, D. C. and Young, D. K. 1970. The influence of deposit-feeding organisms on sediment

stability and community trophic structure. J. Mar. Res. 28: 150-178

Sanders, H. L. 1958. Benthic studies in Buzzards Bay. I. Animal-sediment relationships.

Limnology and Oceanography 3: 245 - 258.

Snelgrove, P. V. R. and C. A. Butman. 1994. Animal-sediment relationships revisited: Cause

versus effect Oceanogr. Mar. Biol. Ann. Rev. 32: 111-177.


Pre-laboratory Report 3, Vertical profiles in Bellingham Bay

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


1: The average salinity of deep water entering Bellingham Bay near the mouth of the bay is 32

psu. The average salinity of surface water exiting Bellingham Bay is 29 psu. The mean flow of

the Nooksack River at this time is 5000 ft3/sec. What is the rate of mean circulation in

Bellingham Bay (in m/sec)?

Rate of average inflow:____________

Rate of average outflow:___________

The area of Bellingham Bay is approximately 40 km2. If the average water depth below the

pycnocline is 85 m, what would be the average bottom-water residence time in Bellingham Bay?

Bottom-water residence time: ________

Show your calculations here:

2: The dissolved oxygen concentration in deep water entering Bellingham Bay is 81 μM. The

lowest dissolved oxygen concentration measured in the center of the bay is 58 μM. If the water

depth is 90 m and the depth of the pycnocline is 5 m, what is the rate of respiration (dissolved

oxygen removal) in the bottom water of Bellingham Bay?

Respiration rate (μmol O2/L/day)_______________

Show your calculations here:


ESCI 322 Lab Report 2 Part 1, Effects of the Nooksack River on biological,

physical and chemical properties of Bellingham Bay

Objectives: Study how the Nooksack River influences Bellingham Bay.

Broader learning objectives: Learn how to collect and process electronic data and seawater

samples using a CTD and rosette, consider processes that influence vertical profiles of seawater

properties, learn how to calculate rates of estuarine circulation and oxygen consumption,

contribute data to a longer-term monitoring program.

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 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 fivefold. First, we’ll examine how freshwater input affects the

biology, chemistry and physics of Bellingham Bay. Second, we’ll observe the distribution of

dissolved oxygen in the water column and consider what processes control DO concentration.

Third, we'll collect samples for chlorophyll so that we can examine the distribution of

phytoplankton biomass relative to Nooksack River input. Fourth, we'll collect nutrient samples

that we will analyze in future lab assignments. Finally, 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:

Salinity and temperature: These properties not only affect biology, but also determine seawater

density, which in turn drives (baroclinic) estuarine circulation.

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

somewhat 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,

from an underwater vantage point, light comes from all directions. The spherical (4pi) sensor

allows accurate measurement of this diffuse light field.

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.

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.

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 that holds six Niskin bottles for collecting

water samples. It will allow us to observe water properties as we lower it into the water. We

will calculate 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 (‰). Temperature is

expressed as degrees Celsius (°C). Other instrumentation on the CTD will measure light,

dissolved oxygen, and seawater fluorescence, which is related to the concentration of chlorophyll

in the water. The six Niskin bottles can be closed to collect water at different depths using a


remote electronic closing mechanism that we will fire as the instrument ascends. We will collect

a sample of surface water at each station and collect water samples from several depths at the

deeper stations. At shallower stations we will collect seawater using buckets. 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).

Water samples: Use a clean bucket to obtain a surface water sample from the shallow stations.

At deeper stations, collect water from the CTD rosette. Close (“fire’) the bottles electronically at

selected depths using the CTD deck unit or software. 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 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.

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. 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.

Next week we will measure chlorophyll on samples we collected this week.

Relationship between temperature, salinity, and water density: Both temperature and salinity

affect seawater density and density () can be calculated 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.

You can 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? (Assess this by using the equation of state to

calculate the density of seawater for different values of salinity and temperature within the

ranges we measured.) How does the Nooksack River affect salinity in Bellingham Bay? Are

hypoxic conditions apparent anywhere in Bellingham Bay? Where is the DO lowest?

Calculation of extinction coefficients: 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 meter depth is given by the extinction (or attenuation)

coefficient k (m-1). The light attenuation coefficient, KD, can calculated from the Secchi depth

(ZSD) according to the formula KD=1.44 /ZSD. Alternatively, the extinction coefficient KD can

also be calculated from vertical profiles of irradiance 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. Consider the following question: How does light

extinction vary with distance from the river, or with fluorescence?

Estimating mean rates of estuarine circulation and water column oxygen consumption: The

mean rate of estuarine circulation (also called residual circulation) can be estimated from

measurements of salinity and flow of the Nooksack River. The USGS measures the Nooksack

River flow just before it enters Bellingham Bay. The data can be accessed at

http://wa.water.usgs.gov/cgi/realtime.data.cgi?basin=nooksack. Click on the Ferndale gauge

station (Stn No. 12213100) to access real-time data. The average rate of water outflow from

Bellingham Bay into the Strait of Georgia can be calculated as follows:

where R is the volumetric rate of river discharge, Si is the salinity of incoming water, S0 is the

salinity of outgoing water, and T0 is the volumetric rate of water outflow (in the same units as R).

In Bellingham Bay, the incoming salinity (Si) is the salinity of deep water that enters the bay in

the south, just east of Eliza Island (station E in Fig. 1). The outgoing salinity (S0) is the average

salinity of surface water leaving the bay (Stns G and F) above the pycnocline.

We can also calculate the average rate of water inflow as follows:

RSS

ST

Oi

i

O

RTTi

0


And, the bottom-water residence time (RT) can be calculated as follows:

where VB is the volume of bottom water. In Bellingham Bay, VB can be calculated as the

product of the area of the bay ABW and the bottom-water depth DBW (distance between the

bottom and the pycnocline). For Bellingham Bay, use ABW = 40 km2. and get DBW from the

CTD profile and station data that we collect.

An estimate of the rate of oxygen consumption in bottom water is the difference in oxygen

concentration between incoming water (deep water at station E) and the lowest oxygen

concentrations measured in the center of the bay divided by the residence time:

Respiration = (Stn E [O2] - minimum [O2]) / RT.

In Puget Sound, oxygen consumption rate range from around 0.2 to 0.8 mmole m-3 d-1 (Barnes

and Colias 1958, Christensen and Packard 1976).

References: Barnes, C. A. and E.E. Collias. 1958. Some considerations of oxygen utilization rates in Puget

Sound. Journal of Marine Research 17:68-8.

Christensen, J.P. and T.T. Packard. 1976. Oxygen utilization and plankton metabolism in a

Washington fjord. Estuarine and Coastal Marine Science 4:339-347.

Gill, A. E. 1982. Atmosphere-Ocean Dynamics, Academic Press, San Diego, CA.

i

B

T

VRT


Figure 1. Station locations for Bellingham Bay survey

Figure 2. Predicted tidal levels in Bellingham Bay, October 21, 2014.


Figure 3. Additional chart of Bellingham Bay


Pre-laboratory Report 4: Seawater fluorescence, chlorophyll, and

phytoplankton biomass

Name: _______________________________

1: Plot the locations of each station in Bellingham Bay that we sampled last week using any

program you like. Plot profiles of temperature, salinity, dissolved oxygen and density (t,

calculated using formula in the provided spreadsheet. Create a contour plot of surface salinity.

2: Read the description of this week’s laboratory assignment and access the Excel spreadsheet

"Chl template" 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.

You filter two 150 ml of seawater samples onto glass fiber filters. (The samples are of surface

and deep water.) After dissolving the samples on the filters in 10 ml of 90% acetone you

measure the samples' fluorescence on a fluorometer, add two drops of 10% HCl, and measure the

fluorescence a second time.

Here are your results

Sample First fluorescence reading Second fluorescence reading

Surface sample 17000 10000

Deep sample 18000 16000

Blank reading 208 203

The K factor (Kx) for the fluorometer is 6.5 x 10-7. F0/Fa = 1.8. Fm = 2.2.

Questions:

What is the chlorophyll concentration in the surface and deep samples?

What is the phaeopigment concentration for both samples?


ESCI 322 Lab Report 2, Part 2, Seawater fluorescence, light, chlorophyll, and

phytoplankton biomass in Bellingham Bay.

Objectives: Measure chlorophyll on samples collected last week.

Broader learning objectives: Learn why chlorophyll is used by biological oceanographers to

estimate phytoplankton biomass. Learn the difference between chlorophyll measurements and

seawater fluorescence. Think about what controls the distribution of phytoplankton biomass in

Bellingham Bay.

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).


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).

Fig. 3. Absorbance spectra of

some commonly occurring

chlorophylls and carotenoids.


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) sonicating 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

(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) 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 <95; the instrument


response is not linear outside this range. You will need to find the correct sensitivity setting for

use with each sample. (If the reading is off-scale on the 1x setting, you will need to dilute your

extract; see step 1, above.) Record sample name, volume of water or culture filtered, volume of

acetone used for extraction, and the fluorescence reading, including the sensitivity setting.

f) Without removing cuvette from fluorometer, add 2 drops of 1 N HCl. Record the fluorescence

after the reading stabilizes. Do not change the sensitivity setting, even if the new reading is <25.

Rinse cuvette well (3x) with 90% acetone to remove any acid.

Step 4: Calculating the chlorophyll concentration

Calculate chlorophyll concentration in each water sample using the following equations (from

Lorenzen, 1966):

Chl a (µg/liter seawater) =)1(

)(0

mf

amx

Fv

dFFvFK

Phaeopigments (µg/liter seawater) =)1(

)(0

mf

ammx

Fv

dFFFvFK

where:

Fo = fluorescence before acidification

Fa = fluorescence after acidification

Fm = maximum acid ratio which can be expected in the absence of pheopigments ≈ 2.2

Kx = calibration factor for a specific sensitivity scale units: [(µg Chl a/ml solvent)/instrument

fluorescence unit]

k1x = 7.12 x 10-4, k3x = 2.40 x 10-4, k10x = 6.43 x 10-5, k30x = 2.48 x 10-5

v = volume of acetone used for extraction (ml)

vf = volume of seawater filtered (liters)

d = extract dilution factor (e.g. if you diluted 1 ml extract by adding it to 4 ml solvent, your

dilution factor would be 5. If no dilution, d = 1).

Note that most of these factors reduce to a constant for a given set of instrument calibration

factors. I will post a spreadsheet containing these formulas for your convenience.

Report:

Address the question of how freshwater input from the Nooksack River affects physical,

chemical and biological properties of Bellingham Bay.

Include the following in your report:

1. Compare chlorophyll-a concentration and seawater fluorescence using linear regression.

Determine the regression equation and the R2 value.

2: For each sampling station, plot chlorophyll and phaeopigment concentrations from the discrete

samples. Also plot σt, light intensity, and fluorescence using units µg Chl-a/liter seawater.

(You'll need to use the linear relationship from (step 1) to convert raw fluorecence units to µg

Chl-a/liter.) Calculate the light attenuation coefficient at each station using both the secchi disk

measurement and the light profile from the spherical light sensor.

3: Consider the following additional questions in your report. How does the Nooksack River

affect salinity, density, and mean circulation? How do chloropigments vary in the vertical and


with distance from the mouth of the Nooksack River? How do the profiles of light, Chl-a and

phaeophorbide-a compare? What might account for these relationships? What is the rate of

bottom water oxygen consumption and how does it compare with measured rates?

References

Jeffrey, S. W., R. F. C. Mantoura, and S. W. Wright, Eds. 1997. Phytoplankton pigments in

oceanography. Monographs on oceanographic methodology. Paris, UNESCO.

Lee, R. E. 1999. Phycology (3rd ed.). Cambridge, Cambridge University Press.

Lorenzen, C. J. 1967. Determination of chlorophyll and phaeo-pigments: spectrophotometric

equations. Limnol. Oceanogr. 12: 343-346.

Parsons, T. R., Y. Maita, and C. M. Lalli. 1984. A manual of chemical and biological methods

for seawater analysis. Oxford, Pergamon.

Thurman, H. V. 1997. Introductory Oceanography (8th ed.) Upper Saddle River, Prentice-Hall.


Pre-laboratory Report 5, Nutrients in Seawater

Name: _______________________________

Read the description of this week’s laboratory assignment and the following questions to be

turned in at the beginning of the lab period. You do not need to type your answers to these


1: What kinds of phytoplankton depend upon silicate as a nutrient?

2. You measure the concentration of a nutrient by means of a colorimetric method using an

absorbance spectrophotometer. Create a standard curve and calculate the concentration of an

unknown sample using the following data.

Standard concentration Absorbance

0 µM 3.1

10 µM 7.4

50 µM 23.1

100 µM 44.2

200 µM 86.0

Absorbance reading of sample with unknown concentration: 76.1


ESCI 322 Lab Report 3: Nutrients in Seawater

Objectives: The objective of this laboratory session is to learn standard techniques for the

analysis of dissolved ammonium and nitrate.

Broader learning objectives: Make a primary standard, a secondary standard, and a standard

curve. Learn precision pipetting. Learn the basics of colorimetric analysis of solutes. Observe

how an autoanalyzer works.

Nitrogen availability often limits productivity of coastal marine systems. Phosphorus can

sometimes be important too, especially in estuaries. Silica availability can limit abundance of

some groups that require silica, such as diatoms. We will obtain data on silicate, phosphate,

nitrate, and ammonium from samples we collected in Bellingham Bay. You will learn to

construct a standard curve for the calculation of nutrient concentrations in each of your samples;

after calculations are done, we will interpret the measured nutrient concentrations by discussing

sources and sinks of these substances. We will also examine the ratios of N and P in Bellingham

Bay and address the question whether concentrations of these nutrients follow Redfield ratios

(discussed below).

Most analytical methods for dissolved nutrient determination are colorimetric. Nutrients react

with reagents, producing colored compounds. The intensity of the color is quantified using a

spectrophotometer set at the appropriate wavelength. Color intensity is linearly related to the

amount of nutrient present. A standard curve allows us to relate the color intensity to the nutrient

concentration in the sample.

Part I: Nitrate (+ nitrite) and Ammonia

Concentrations of nutrients in marine waters are controlled by both the rate of nutrient input and

the rate of uptake by organisms. Phytoplankton take up nutrients in a consistent ratio, known as

the Redfield ratio. The Redfield ratio is the ratio in atoms of carbon, nitrogen, and phosphorus

assimilated by phytoplankton. This ratio is 106:16:1 (C:N:P). In addition, silicate often has a

1:1 ratio with phosphorus, depending upon the composition of the phytoplankton community.

Because concentrations of nutrients are so strongly tied to biological uptake, concentrations of

dissolved nutrients often follow this ratio. This week, by measuring nitrate + nitrite and

ammonium, the main inorganic nitrogen species in seawater, we can calculate the concentration

of total dissolved inorganic nitrogen ([DIN] = [NO3- + NO2

- + NH4+]) and compare it to the

concentrations of P and Si d (to be measured the following week) to determine whether

concentrations of nutrients in Bellingham Bay follow the Redfield ratio.

Analysis of dissolved Ammonium Outline of method (from Parsons et al. 1984): Seawater is treated in an alkaline citrate medium with

sodium hypochlorite and phenol in the presence of sodium nitroprusside which acts as a catalyser. The

blue indophenol color formed with ammonia is measured spectrophotometrically.

Procedures: Wear gloves and goggles. Make sure the spectrophotometer is turned on and set to a

wavelength of 640 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.

Reagents: Reagents have also been made ahead of time. This procedure uses three: phenol solution,

sodium nitroferricyanide, alkaline reagent, sodium hypochlorite, oxidizing solution. These are

hazardous. Use caution and wear gloves and eye protection.

Phenol solution: 20g phenol in 200 ml 95% v/v ethanol

Sodium nitroferricyanide solution: 1 g sodium nitroferricyanide, in 200 ml D.I. H2O (Store in

dark glass bottle. Stable for one month.)

Alkaline reagent: 100 g sodium citrate and 5 g NaOH in 500 ml DI. H2O

Sodium hypchlorite solution: CloroxTM bleach (with no whiteners or additives)

Prepare oxidizing reagent: 100 ml alkaline reagent and 25 ml sodium hypochlorite solution (Chlorox).

Keep stoppered and prepare fresh daily. Ammonia standards Primary standards – 0.100 g ammonium sulphate (NH4)2SO4 in 1 L Nanopure water. Secondary standards – to be prepared from primary standards.

2° STD (M) L 1° STD/50 ml H2O 0.00 0 0.75 25 1.50 50 2.25 75 3.00 100 5.50 150

Quality control standard:

1.50 Procedure:

1. Pipette 5 ml aliquots of each secondary standard (2x for high standard), D.I. water blanks (2x), quality control standards (2x), and each sample (1x) into culture tubes

2. In fume hood, add 0.2 ml phenol, vortex, 0.2 ml nitroferricyanide, vortex, 0.5 ml oxidizing solution, vortex, and store in the dark for 1 h.

3. Vortex each sample and measure the absorbance at a wavelength of 640 nm in a spectrophotometer.

Calculations: The [NH4] of each sample is calculated as: [NH4] (M) = Sample absorbance - blank absorbance

Slope of standard curve (absorbance/M)

Analysis of dissolved nitrate

We will use an autoanalyser (Westco Smartchem) to analyze our samples for nitrate and nitrite. The

autoanalyser is perhaps the most common technology used for nutrient analysis in oceanography. This

allows large numbers of samples to be processed rapidly and high quality data can be produced by this

method.

Outline of method (from Parsons et al. 1984): Nitrate in seawater is reduced almost quantitatively to

nitrite when a sample is run through a column containing cadmium coated with metallic copper. The

nitrite produced is determined by diazotizing with sulfanilamide and coupling with N-(1-naphthyl)-


ethylenediamine (NED) to form a highly colored azo dye which can be measured spectrophotometrically.

Any nitrite initially present in the sample must be corrected for, otherwise the result is reported as nitrate

+ nitrite.

Nitrate standards Primary standards – 10mM potassium nitrate solution. Dissolve 0.76 g potassium nitrate in 250 ml Nanopure water Refrigerate. (I'll prepare the nitrate standard ahead of time.) Reagents have also been made ahead of time. This procedure uses four: imidazole buffer, cupric sulfate

(0.01 M and 2%), sulfanilamide, and N-(1-naphthyl)-ethylenediamine (NED)

Imidazole buffer (0.1 M): Dissolve 3.40 g imidazole in 500 ml H2O. Dissolve imidazole in 400 ml H2O

in a beaker with a pH electrode and stirrer. Adjust solution to pH 7.0 with HCl. Transfer to 500 ml vol.

flask and dilute to mark. Refrigerate.

Cupric sulfate (0.01M): Dissolve 0.635 g cupric sulfate in 250 ml H2O. Stable.

Cupric sulfate (2% w/v): Dissolve 5 g cupric sulfate in 250 ml H2O. Stable.

Sulfanilamide: Slowly add 50 ml conc HCl to 350 ml H2O in 500 ml volumetric flask.

Dissolve 5 g sulfanilamide and dilute to mark. Stable.

NED: Dissolve 0.5 g NED in 500 ml H2O in volumetric flask. Refrigerate in brown bottle.

Pipette calibration Effects of temperature on fresh water density Temperature (°C) Density (g/ml)

10 0.9993

15 0.9977

20 0.9951

25 0.9914

27 0.9897

28 0.9888

30 0.9869

Prelaboratory report 6, Nutrients in seawater, due next week:

Calculate the concentrations of ammonium and nitrate (+ nitrite) in the samples we collected

from Bellingham Bay and Burrows Bay. Plot these data as profiles.


ESCI 322 Lab Report 3, Nutrients in Seawater Part II - Phosphate and silicate

Objectives: Measure silicate and phosphate in our samples from Bellingham Bay.

Broader learning objectives: Learn how to create a truly zero blank using the reverse-addition

method. Learn how to interpret data on nutrients and other "non-conservative" tracers by

plotting the data versus salinity.

Analysis of dissolved silicate

Silicate is an important nutrient for diatoms and can be a limiting nutrient when nitrate and

phosphate are in high concentrations. Low concentrations of silicate relative to other nutrients

has been linked to shifts in phytoplankton species composition from diatoms to flagellates and

has been implicated in the formation of harmful dinoflagellate blooms.

Outline of method:

The seawater sample is allowed to react with molybdate under conditions which result in the

formation of silicomolybdate. A reducing solution, containing metol and oxalic acid, is then

added, which reduces the silicomolybdate complex to give a blue color and simultaneously

decomposes any phosphomolybdate and aresenomolybdate. The resulting extinction is measured

in a glass cuvette using a spectrophotometer at a wavelength of 810 nm.

Procedures:

Make sure the spectrophotometer is turned on and set to a wavelength of 810 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.

Use plastic (polypropylene) for everything. Glass is made of SiO2, the compound we are trying

to measure. In fact, the Earth's crust is ~60% SiO2. Consequently, contamination will be your

biggest obstacle to measuring silicate. Triple-rinse all labware just before use. Wear clean

gloves when handling samples, standards, and reagents. Keep your sample containers, standards,

and reagents free from dust.

Standards: To save time, we will use previously-made standards (5, 10, 20, 30, 40, and 50M

SIO4). Ordinarily, these would be made by dissolving Na2SiF6 in DIW.

Reagents: Reagents have also been made ahead of time. This procedure uses three: Metol-

sulfite reagent, oxalic acid, and sulfuric acid. These are hazardous. Use caution and wear

gloves and eye protection.

Procedure:

4. Pipette 2.5 ml of sample or standard into a clean 10-ml polypropylene centrifuge tube. 5. Add 1.0 ml of acidified ammonium molybdate reagent to each sample and standard. But

do not add this reagent to the blank [0M Si(OH)4]. Cap tightly and shake once or twice. The sample will now be at pH 1.0 - 1.5, and the following reaction will go to completion in about 10 minutes:


Si(OH)4 + 12 MoO4= + 24 H+ -> 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.

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 1 sample 10 samples 20 samples 50 samples 100 samples

Metol-sulfite 0.5 ml 5 ml 10 ml 25 ml 50 ml

Oxalic Acid 0.3 ml 3 ml 6 ml 15 ml 30 ml

H2SO4 0.3 ml 3 ml 6 ml 15 ml 30 ml

DIW 0.4 ml 4 ml 8 ml 20 ml 40 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. This means the reverse-addition blank contains all the reagents and other colored compounds, but none of the blue-colored silica complex.

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-, HPO4

2-, 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.

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 the 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 or beaker several times with a few ml of the

secondary standard solution. Using a volumetric pipette, carefully add the secondary standard

solution from the beaker 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) ml 2° STD / 100 ml H2O

0.0 0.00

0.6 0.50

1.8 1.50

3.0 2.50

4.2 3.50

Place six glass culture tubes in a tube rack (two for the blank and high standard) and record their

positions. Also add tubes for the seawater samples and record their positions.

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 0.1 ml Ammonium molybdate, 0.25 ml Sulfuric acid, 0.1 ml Ascorbic acid,

and 0.05 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 10 samples 20 samples 50 samples

Ammonium molybdate 1.0 ml 2.0 ml 5.0 ml

Sulfuric acid 2.5 ml 5.0 ml 12.5 ml

Ascorbic acid 1.0 ml 2.0 ml 5.0 ml

K-antimonyl tartrate 0.5 ml 1.0 ml 2.5 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 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 within the bay without

reference to specific locations. We can even infer whether biological or chemical processes produce or

consume nutrients within Bellingham Bay as well. This method is described next.


Understanding relationships between salinity and nutrient concentrations

Mixing theory: A non-reactive solute (sometimes referred to as a conservative tracer) supplied by a river

will be diluted by seawater. If dilution is the only process that determines the concentration of the

ttracer in an estuary, the relationship between tracer concentration and salinity will be linear. This also

holds for a tracer with a seawater input.

Figure 1. Theoretical

relationship between a

conservative solute and

salinity for solutes with (A)

a river source and (B) a

marine source.

A non-conservative tracer is one that has a source or sink within the estuary. In this case, the

relationship between tracer concentration and salinity will be non-linear.

Figure 2. Theoretical

relationships between non-

conservative 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.

Salinity

Tra

cer

co

nce

ntr

ation

Salinity

Tra

cer

co

nce

ntr

ation

River source Marine sourceA B

Salinity

Tra

cer

co

nce

ntr

ation

Salinity

Tra

cer

co

nce

ntr

ation

River source Marine sourceA B

Salinity

Tra

cer

concentr

ation

Salinity

A B

Estuarine source

Estuarine sinkSalinity

Tra

cer

concentr

ation

Salinity

A B

Estuarine source

Estuarine sink


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. 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. Denitrification in marine sediments can

reduce N:P whereas sorption of P onto sediments could

increase the ratio. Also, preferential uptake of N versus P in

the marine waters entering the bay could reduce N:P.

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 and deep water inflow from the Strait of Georgia/Strait of Juan de Fuca. 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 using

the same methods as before: Ti = R[SO/(Si-SO)] = TO - R

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 (for each nutrient,

include all data from all stations), examine vertical profiles, and compare your data with theoretical

expectations. Present your nutrient budget as a table in your report including total inputs and % inputs.

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.

Salinity

DIN

:DIP

ratio

16

8

0

Redfield ratio

N < P:

Denitrification

In sediments?

N < P:

Fertilizer input?

N > P:

Nitrogen fixation?

N > P:

Phosphate storage

In sediments?

Salinity

DIN

:DIP

ratio

16

8

0

Redfield ratio

N < P:

Denitrification

In sediments?

N < P:

Fertilizer input?

N > P:

Nitrogen fixation?

N > P:

Phosphate storage

In sediments?


Pre-laboratory Report 7, 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 Initial chlorophyll Final chlorophyll Net growth

(fraction seawater) concentration (p0) concentration (p) 1/t * ln(p/p0)

1.00 30 38.5

0.75 22 53.3

0.50 11 47.1

0.25 3.0 29

0.05 0.15 4.0

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: ________________


ESCI 322 Lab Report 4: Phytoplankton population growth and grazing

Objectives: Conduct a dilution experiment for estimating rates of population growth & grazing.

Broader learning objectives: Understand the mathematics and units of population growth and

grazing. Think about the role of grazers in controlling rates of phytoplankton growth.

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

follows: pdt

dp , where p is the concentration of phytoplankton and µ is the per-capita growth

rate. If z is the microzooplankton concentration and χ is the phytoplankton-zooplankton

encounter rate, you can add grazing to this equation as follows: zppdt

dp . Now, consider

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

growth model, you can write it as gDppdt

dp , where g is the per capita mortality rate of

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

gives the following formula: gDp

p

t

0

ln1

. The first term is a measure of phytoplankton

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

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 Number of Light Sampling

Seawater replicates level time

100% 4 NA initial

100% 2 100% final

100% 2 10% final

50% 3 NA initial

50% 1 100% final

10% 4 NA initial

10% 2 100% final

10% 2 10% 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.

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.

Dilution experiments

Dilution factor (fraction SW)

Net

Gro

wth

Ra

te (

per

tim

e)

1

Decreasing # of Grazers

Y-intercept= “infinite dilution”

GROWTH RATE

Slope = linear relationship

with dilutions

HERBIVORY RATE

Growth equation: dp/dt = p – zp

dp/dt = p – zDp = p – gDp

Solution: 1/t ln(p(t)/p0) = – gD

0

Dilution experiments

Dilution factor (fraction SW)

Net

Gro

wth

Ra

te (

per

tim

e)

1

Decreasing # of Grazers

Y-intercept= “infinite dilution”

GROWTH RATE

Slope = linear relationship

with dilutions

HERBIVORY RATE

Growth equation: dp/dt = p – zp

dp/dt = p – zDp = p – gDp

Solution: 1/t ln(p(t)/p0) = – gD

0


ESCI 322 Lab Report 4: Phytoplankton population growth and grazing

Week 2: Sample processing and analysis

Objective: Measure chlorophyll as a marker for phytoplankton biomass and estimate population

growth and grazing from the dilution experiment results. Identify phytoplankon groups

Broader learning objective: Learn to identify and enumerate phytoplankton. Also, observe

ciliate grazers from our experiment.

Reconsider the population growth equation that forms the basis of our dilution experiment:

gDp

p

t

0

ln1

. Note that ln(p/p0) is unitless. It depends on the ratio of p/p0 (the ratio of

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.

Review of chlorophyll measurement 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

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 <95; the instrument

response is not linear outside this range. You will need to find the correct sensitivity setting for

use with each sample. (If the reading is off-scale on the 1x setting, you will need to dilute your

extract; see step 1, above.) Record sample name, volume of water or culture filtered, volume of

acetone used for extraction, and the fluorescence reading, including the sensitivity setting.

f) Without removing cuvette from fluorometer, add 2 drops of 1 N HCl. Record the fluorescence

after the reading stabilizes. Do not change the sensitivity setting, even if the new reading is <25.

Rinse cuvette well (3x) with 90% acetone to remove any acid.

Step 4: Calculating the chlorophyll concentration

Calculate chlorophyll concentration in each water sample using the following equations (from

Lorenzen, 1966):

Chl a (µg/liter seawater) =)1(

)(0

mf

amx

Fv

dFFvFK

Phaeopigments (µg/liter seawater) =)1(

)(0

mf

ammx

Fv

dFFFvFK

where:

Fo = fluorescence before acidification

Fa = fluorescence after acidification

Fm = maximum acid ratio which can be expected in the absence of pheopigments ≈ 2.2

Kx = calibration factor for a specific sensitivity scale units: [(µg Chl a/ml solvent)/instrument

fluorescence unit]

k1x = 7.12 x 10-4, k3x = 2.40 x 10-4, k10x = 6.43 x 10-5, k30x = 2.48 x 10-5

v = volume of acetone used for extraction (ml)

vf = volume of seawater filtered (liters)

d = extract dilution factor (e.g. if you diluted 1 ml extract by adding it to 4 ml solvent, your

dilution factor would be 5. If no dilution, d = 1).

Note that most of these factors reduce to a constant for a given set of instrument calibration

factors. I will post a spreadsheet containing these formulas for your convenience.

Determining cell densities:

Determine the density of phytoplankton and microzooplankton in your samples by counting them

with a Sedgwick Rafter counting slide. This slide holds exactly 1 ml of sample. The 10-ml


samples were preserved with Lugol’s iodine solution in 15-ml centrifuge tubes.

Before counting the samples, perform a tenfold cell concentration step. Centrifuge the samples

and, using a glass pasture pipette, carefully remove the top 9 ml of solution leaving just under 1

ml in the tube. Then, transfer the remaining solution into the Sedgwick rafter cell. Do this by

placing the slide cover on top of the cell at an angle leaving two small triangular openings on

each side of the cell. Dispense your sample into one of these openings and fill the cell. If the

sample does not fill the entire cell, add some distilled water. Once the cell is filled, twist the

cover slip to enclose the sample. To count the sample, place the counting cell on a compound

microscope. Start on one corner of the slide and count “lanes” that cross the entire cell. At the

end of each lane, move the slide the width of one field of view and count the next adjacent lane.

Repeat until you’ve covered the entire slide. Record the number and types of plankton you

observe. Count approximately 300 individual plankters in each sample. Calculate the percent

abundance of each type of alga.

Report

Calculate the per-capita growth and grazing rates for each treatment. Your report should address

the following questions: How important is grazing in controlling phytoplankton growth? How

might the relationship between light intensity, growth and grazing influence the vertical

distribution of phytoplankton biomass?

Reference

Lorenzen, C. J. 1966. Determination of chlorophyll and pheo-pigments: spectrophotometric

equations. Limnol. Oceanogr. 12: 343-346.

envr 322 oceanography laboratory

Documents