1. science
2. biology
3. marine biology

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 12, 2012
ECSI 322 – Oceanography Laboratory - Manual
1
ESCI 322 - Introduction to Oceanography Laboratory
Course Syllabus
Instructor: David Shull
Office: ES 445, ext. 3690
Email: [email protected]
OBJECTIVES:
1.
Acquire first-hand experience with oceanographic research methods
2.
Become familiar with marine organisms and coastal processes in Puget Sound
3.
Learn field and laboratory techniques
4.
Practice scientific writing (writing proficiency course - WP3)
5.
Use oceanographic methods to address local marine environmental problems.
COURSE OVERVIEW:
We will use oceanographic methods to study ecosystem functions and environmental problems
in the marine waters near Bellingham. Students will write reports addressing these issues. Many
of our "labs" will take place in the field, either on a boat or at the Shannon Point Marine Center.
EVALUATION GUIDELINES:
Assignments:
Students will be evaluated based on the completion of several lab reports, a few smaller lab
assignments, and “pre-laboratory” assignments. Each assignment will be typed, although figures
and graphs may be drawn by hand. The reports will be typed and will follow the standard
scientific format; abstract, introduction, methods, results, discussion, references. For some of the
reports, a draft of a portion of the results section will be handed in first for evaluation and
comments. Comments on the draft of the results section are intended to aid students in the
completion of the final reports. Both the results-section draft and the final reports must be turned
in on time to receive full credit. Late reports will receive a 10% grade reduction each day until
the report is turned in. Details on the format of the major reports are given in the section of the
syllabus entitled "Laboratory Report Format". Read this section carefully before you begin. The
smaller assignments will be completed in question-and-answer format. All reports should be
typed, double-spaced, and checked for correct grammar and spelling. You should read through
the assignment, make notes, and think through the organization of all your responses before
writing. Pre-laboratory assignments needn’t be typed but must be handed in at the beginning of
each laboratory period to receive credit.
Contributions of assignments to final grade:
Full lab reports
Draft Reports
Pre-lab assignments
80%
10%
10%
Approximate grading scale:
93-100 A
90-92 A88-89 B+
73-77 C
69-72C67-68 D+
80-82 B57-60 D-
83-87 B
61-66 D
ECSI 322 – Oceanography Laboratory - Manual
78-79 C+
0-56 F
2
Policy for late assignments and reports:
Final reports will lose 10% of the final grade for each late day until turned in. (Reports received
after 5 PM will be picked up the next day.) Pre-labs and draft reports will not be accepted late.
Lab schedule:
Date Topic Location Draft 2‐Oct Waves ES 60 9‐Oct Habitat utilization by rockfish I SPMC* 16‐Oct Habitat utilization by rockfish II WWU 23‐Oct Oxygen, chlorophyll, and circulation in Bellingham Bay SPMC* 30‐Oct Chlorophyll and phytoplankton biomass SPMC 6‐Nov Nutrients in Seawater I ES 331 13‐Nov Nutrients in Seawater II ES 331 20‐Nov Measuring phytoplankton growth and grazing I SPMC 27‐Nov Measuring phytoplankton growth and grazing II SPMC 4‐Dec Zoea required Final Report 9‐Oct 16‐Oct 23‐Oct 30‐Oct 6‐Nov + ROV + CTD 13‐Nov 20‐Nov 27‐Nov 4‐Dec Dead week: Meet to discuss final paper due on Friday *On these days we will be outside for part or all of the lab session. Dress for cold, rain and wind
LAB REPORT FORMAT
A laboratory report is a document in the form of a scientific paper. Scientific papers are the
means by which scientists communicate their research findings to one another. Writing up and
publishing research results is just as important as conducting research in the first place, for if
results are not made available to others, they are of little value. For ease of communication, there
is a generally accepted format for writing up scientific data that we will follow in this course.
Mastery of scientific writing skills is a vital component of becoming a scientist. Scientific papers
have 7 sections: Title, Abstract, Introduction, Materials and Methods, Results, Discussion, and
References. These headings should be placed at the beginning of each section in your report
(except “Title”).
Brief descriptions of the seven sections follow.
Title
The title should be a self-contained explanation of the information presented in your paper. It
must contain enough detail to be informative, without being so long as to be incomprehensible.
Avoid vagueness at all cost.
Too short: Vertical profiling
Too vague: Oceanography laboratory report #1
Too long: A student investigation of the effects of the Nooksack River on the vertical and
horizontal distribution of temperature, salinity, density, nutrients, and dissolved oxygen in
Bellingham, Bay, WA, in November, 2010.
ECSI 322 – Oceanography Laboratory - Manual
3
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, the major results, and inferences drawn from
those findings. If the development of new methods is an important part of the paper, they should
be described in the abstract as well. References are not typically cited in the abstract.
Introduction
The introduction sets the stage for the presentation of your research results and their
interpretation. It must include some background information, to bring the reader up to speed on
the general issues, some specifics, to acquaint the reader with your particular investigation, and
the questions or hypotheses that you will be addressing with the data. Effective introductions are
usually short (several paragraphs).
Background information: What is the general problem that is being studied? What is your
specific approach to that problem? If there is relevant background literature (other important
scientific papers that set the stage for your work), this is the place to briefly summarize their
findings and importance. However, the introduction should not be a literature review.
Specifics: This section will vary depending on the type of research you are presenting. In an
environmental study, you should let the reader know where and when you were working, and
what the environment was like in a general way. If you were presenting the results of an
experiment with organisms, you should describe the species used and the general approach. Try
to develop a logical flow from the “big picture” of background information to the specifics of the
system you studied.
Research questions: End your introduction with a concise summary of the research questions,
hypotheses or goals. You will come back to these in the concluding paragraph of your
discussion.
Materials and Methods
The materials and methods section describes how, when, where and what you did. Describe the
procedures, equipment, and experimental set-ups in enough detail that the work could be
repeated by another scientist, but without extraneous detail. List the methods or procedures
chronologically (i.e. in the order in which you did them). Be sure to include information about
numbers of replicate treatments or observations, types of instruments and equipment used, etc. If
statistical analyses were performed, state the statistical methods used and the data to which they
were applied. Use the past tense.
Methods that are already published can be referred to with a reference; only deviations from the
published method need to be described in your paper. (“Nitrate was measured by the method of
Parsons et al. (1984), except that reagent additions were scaled to a sample volume of 5 ml.”)
You can reference lab handouts, but be sure they are cited in the reference section. You do not
need to explain things that are not necessary for understanding or repeating the work (“The
ECSI 322 – Oceanography Laboratory - Manual
4
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.)
Figures and Tables: Each figure (map, diagram, or graph) and table in a results section should
have a “reason for being.” Don’t present data just because you collected it; present data only if it
you refer to it in the text and it contributes to the story that you are telling in your paper. In
general, figures are plotted with the independent variable on the x axis. Vertical profiles in
oceanography have a special format in which the independent variable (depth is plotted on the yaxis. (We'll discuss this more in class.) Each table and figure should be readily understandable
without reference to the text. Each should have a consecutive number (Fig. 1, 2, 3…); tables are
numbered separately from figures. Finally, each figure and table should have a caption
(sometimes called a ‘legend’) that concisely describes the content (e.g.: Fig. 2. Average rates of
respiration (±1 s.d.) over time for the anemone Anthopleura elegantissima at 10ºC.)
Text: The text of the results section should weave the data presented in figures and tables into a
coherent story. Prepare the figures and tables first, and then write the text. Do not reiterate all the
details of the data; rather, tell a story that describes the major features and any clear trends or
patterns. Refer directly to the appropriate figures and tables, by number, in your text. The first
figure referred to should be Fig. 1, the second Fig. 2, etc. Keep the writing simple and direct.
Don't use the word Figure or Table as the subject of a sentence, e.g., Figure 1 shows the
locations of the sampling sites, because this ruins the narrative style. Instead, tell the story and
refer to figures in parentheses.
Examples:
Not good: The graph of temperature versus depth looks linear near the bottom.
Still not so good: Fig. 1 shows that temperature was constant with depth near the bottom.
Better: Temperature was constant throughout the bottom 5 m of the water column (Fig. 1).
Incorrect: A paired t-test showed that the respiration measurements were significantly different
at the 95% confidence level.
Good: Growth rates in the anemones were higher at 12ºC than at 10ºC (t-test, t[8]=2.9, p = 0.02).
Discussion
In the discussion you interpret your results: tell the reader what they mean and why they are
important. In this section you should answer “why?” and “how?” questions about your data. For
example, why was the temperature different at the top relative to the bottom of the water
column? Why did the respiration rate of the anemones increase with temperature? Here reader
ECSI 322 – Oceanography Laboratory - Manual
5
should discover the answers to your original questions or hypotheses as set forth in the
introduction. The discussion section is also where you compare your findings to those of others,
as reported in the scientific literature. You may also want or need to discuss short-comings in
your methods, or the need for further testing. This latter should not, however, be the main focus
of the section. The discussion section should be a general synthesis of your findings and their
importance. Do not restate your results. The key word is interpretation. This section is usually
the hardest to write; think it through carefully and prepare an outline before you begin. One
effective technique is to start the section with your strongest or most important finding.
References
This section is an alphabetical listing, by first author’s last name, of the references cited in your
paper. There are two ways to cite a paper in your text: Several other species of anemone are
known to have respiration rates that increase with temperature (Matthews, 1993; Smith and
Wesson, 1998). Our findings of lower respiration rate at lower temperatures agreed with those
of Matthews (1993). If there are more than two authors, use the term et al. (an abbreviation of et
alias, “and others”) after the name of the first author: Michaels et al. (1994) or (Michaels et al.,
1994). Journals have their own specified format for listing references, which should be followed
when submitting a paper to that journal. For our purposes, you can use the format below.
Journal article:
Name(s) of author(s). Date. Title of article. Title of journal (may be abbreviated). Vol #: pages.
Example:
Lenington, S. 1979. Effect of holy water on the growth of radish plants. Psychological Reports
45: 381-382.
Book:
Name(s) of authors. Date. Title of book. Publisher, Location (city) of publisher, # of pages in
entire book.
Example:
Povinelli, D. J. 2000. Folk Physics for Apes: The Chimpanzee's Theory of How the World
Works. Oxford University Press, New York, 391 pp.
Chapter from a book in which each chapter has a different author and the book has an editor:
Name(s) of authors. Date. Title of chapter. In: editor(s), Book Title. Publisher, Location (city) of
publisher, pages.
Example:
Jumars, P. A. 1993. Gourmands of mud: Diet selection in marine deposit feeders. In: R.N.
Hughes (Ed.), Mechanisms of Diet Choice. Blackwell Scientific Publishers, Oxford. pp.
124-156.
ECSI 322 – Oceanography Laboratory - Manual
6
ESCI 322 Lab Assignment : Waves and Coastal Processes
During this laboratory session we will study some of the properties of waves. We will create our
own experiments in wave tanks, altering variables known to affect wave properties (see below),
to explore the mathematical relationships among those properties. We will also set up an
experiment to simulate the effects of waves on coastlines.
Terminology:
-Wave height (H) is the vertical change in height between the wave crest and the wave trough.
-Wave amplitude (A) is one-half the wave height.
-Wavelength (L) is the distance between two successive peaks or troughs.
-Steepness is wave height divided by wavelength (H/L) (note this is not the same as the slope
between a wave crest and its adjacent trough).
-Wave period (T) is the time it takes for two successive peaks (or troughs) to pass a fixed point.
-Wave frequency (f) is the number of peaks (or troughs) which pass a fixed point per second.
Review of wave relationships:
Wave period is the inverse of wave frequency: T = 1/f
Wave speed (termed celerity) can be related to wavelength and period according to the general
formula: C = L/T.
These formulas hold for all waves. For gravity waves at the water surface, the following
equation can be used to calculate wave speed from wavelength and water depth.
1
 gL
 2d   2
C  
tanh
  ,
 L 
 2
where d is water depth (below mean surface level), and g is the gravitational acceleration (9.81
m/sec2). We will test this theoretical model in part two of our lab assignment. This equation
reduces to simpler forms for deep-water and shallow-water waves.
Deep-water waves: water depth (d) is greater than L/2 and C = gL 2 (m/s). Since g is a
constant, this formula reduces to C= 1.25 L (m/s). Note that L, the wavelength, is the only
variable affecting wave speed for deep water waves. Since L, T and C are related, the equation
for deep water waves can be rewritten as C = 1.56T (m/s).
Shallow-water waves: water depth (d) is less than L/20 and C = gd = 3.13 d (m/s). Note that
d, the water depth, is the only variable affecting wave speed for shallow-water waves.
Intermediate waves: if water depth is < L/2 and >L/20, the more complex formula given above
must be used to calculate wave speed.
Affects of bottom topography:
Refraction: Because wave speed for shallow-water waves is a positive function of water depth,
waves slow as they approach the shoreline. Wave period remains constant so that a decrease in
wave speed reduces wavelength. Parallel-crested waves approaching the shoreline at an angle,
therefore, will refract, bending to become more parallel to the shore before they break. Bottom
topography around bays and headlands will result in refraction patterns causing variation in the
spatial distribution of wave energy, sediment erosion, and sediment deposition along the shore.
ECSI 322 – Oceanography Laboratory - Manual
7
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?
ECSI 322 – Oceanography Laboratory - Manual
8
Place a floating object in the tank near the shoreline. Can you observe longshore transport?
Can you observe areas of erosion or deposition? Let the wave tank run while we move on to
experiment two. After 1 h, repeat the shoreline measurements in the large tank and identify
regions of erosion, deposition, and areas that are stable. Attempt to fix any problems with your
property by dredging, adding a breakwater or groin, by beach nourishment, or by beach
armoring.
6: Let the tank run for another hour and then repeat your drawing and observations.
Results
7: Which of the development projects appear to be a success?
8: Which appear to be failures?
9: Why?
Relationships between C, L, T, f, H, and d:
These experiments will be conducted in the narrow wave channel. Turn on the wave generator
and adjust the wave frequency by turning the small knob on the control box. Adjust the wave
height by opening or closing the valve on the compressed air tank.
Assignment
Measure C, L, T, f, H, or d for waves of different heights and frequencies. Use these
measurements to test the following relationships for transitional waves:
Wave speed:
Properties of breaking waves:
 gL
 2d
tanh 
C  
 L
 2
1
 2
 

H/L = 1/12 (= 0.083) H/D = 0.8
Measuring wave velocities and wave lengths is more difficult than it seems. We will use simple
submerged pressure sensors attached to an oscilloscope to measure wave period (T) and speed
(C). We will then calculate wavelength from the relationship L = CT. An oscilloscope displays
a graph of an electrical signal. It shows
how electrical signals vary over time; the
vertical axis represents voltage and the
horizontal axis represents time. As a wave
passes over the submerged pressure sensor,
the pressure increases according to the
formula P=ρgh, where P is pressure, ρ is
fluid density, and h is the height of the
water above the sensor. The pressure
sensors send a signal (a change in voltage)
which is recorded on the oscilloscope.
10: Make two plots: (a) Measured wave velocity vs. water depth, and (b) measured wave velocity
versus wavelength. Plot the measured wave velocities and the velocities predicted by the
formula on the same graph.
ECSI 322 – Oceanography Laboratory - Manual
9
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
ECSI 322 – Oceanography Laboratory - Manual
Wavelength
10
Assignment summary
1: Patterns of erosion and deposition due to wave action (large wave tank)
- Draw developed shoreline before and after period of wave inundation
- State how you expected the beach to change
- Describe how it actually changed
- Which development projects worked and which did not? Why?
2: Testing the wave velocity formula (wave channel)
- Make two plots: Measured wave velocity vs. water depth, and measured wave
velocity versus wavelength. Plot the measured wave velocities and the velocities
predicted by the formula on the same graph.
- Discuss whether the predicted wave velocities matched the measured velocities, and
try to explain any differences you observe.
3: Effects of breaking waves (wave channel)
- Draw two beach profiles (before and after wave inundation)
- Calculate the ratios H/L and H/D at the location where the waves broke
- Compare the measured ratios with the critical ratios for wave breaking
- Which controls wave breaking in the wave channel?
ECSI 322 – Oceanography Laboratory - Manual
11
Pre-laboratory Report 1, Rockfish Habitat Utilization
Name: _______________________________
Read the description of this week’s laboratory assignment and answer the following questions.
You are watching video of the bottom collected by an ROV. In the center of the images you are
viewing, two glowing points from laser beams are visible. The laser beams on the ROV are 10
cm apart. You measure the distance between the laser dots on the video screen and determine
that the average distance between them on the video screen is 3 cm. The width of the image on
the screen is 15 cm. You count six fish as the ROV traverses 50 m of bottom.
Questions:
What is the actual width of the video of the bottom in cm?
What is the area of the transect in m2?
What is the fish density (number per square meter) in the transect?
ECSI 322 – Oceanography Laboratory - Manual
12
ESCI 322 Lab Report 1: Benthic habitat utilization
Demersal fish and other bottom-dwelling organisms are often associated with particular types of
bottom habitat. These organisms might seek out these habitat types because they provide food or
refuge from predators. The larval stages of these organisms could be passively deposited in
certain habitats or they might actively choose a particular bottom type for settlement.
Alternatively, adults might migrate to these areas. Regardless of the reason behind these
associations, habitat is clearly important to organisms and proper management of marine
resources requires that appropriate habitats are preserved along with species of interest. Habitat
preservation seems particularly important for rockfish in Puget Sound and the Strait of Georgia.
Rockfish populations have been declining in recent decades and many advocate for the
establishment of marine protected areas in Puget Sound to protect rockfish and their habitat
(Palsson et al. 2009). But, what kinds of habitat should be protected?
The purpose of this week’s and next week’s labs is to document patterns of habitat utilization by
rockfish in Whatcom County (or Skagit) using a remotely operated vehicle, or ROV. The ROV
is essentially a submersible video camera with propellers. It is tethered to the surface and
controlled using a joy stick. We will deploy the ROV at several locations either along the southeastern shore of Lummi Island or in Burrows Bay where there exists a gradient in bottom type.
We will deploy the ROV in a manner similar to that depicted in figure 1.
Figure 1. ROV deployment and video collection
plan (from Auster et al. 1997).
Our survey will have two components. For the first,
we will determine the bottom slope using the R/V
Zoea’s GPS to determine our horizontal position
(distance offshore) at several locations and the depth
sounder to determine the water depth at the same
locations. We will then be able to calculate bottom
slope from the plot of bottom depth versus distance
offshore. For the second component, we will drop the
ROV at the deepest part of the sloping bottom and fly
the ROV upslope, toward shore. We will direct the
video camera toward the bottom and record fish and
other organisms within the field of view of the camera.
The ROV is equipped with two parallel lasers, 10 cm
apart, which serve as a horizontal distance scale for the
video. The lasers are set at a known angle from the
horizontal so that the vertical distance scale can also
be determined. To simplify the scale conversions, the
camera tilt should be adjusted so that the laser dots are
centered within the video field of view (Fig. 2). If L is
the distance between the two lasers, and l is the
distance between the two laser dots on the video
screen, and a is the width of the video screen you are
ECSI 322 – Oceanography Laboratory - Manual
13
viewing, then the actual width of the transect, A, is equal to (L/l)a. (If we wanted to know the
other horizontal dimension of the bottom video, B we could calculate it too. If θ is the angle
between the laser and the horizontal, and b is the height of the video frame, then the actual height
of the frame, B, equals (L/l)b/sin(θ).) If we know the length of the transect and its width, we can
calculate the area surveyed. Rockfish density is the number of rockfish per unit area. For our
study, we will determine the width of the transect by calculating A as described above using the
distance between the lasers. We’ll calculate the total transect length from the change in water
depth observed during the video transect and the bottom slope.
camera
B
Ll
θ
laser
A
H
θ
H’’
Figure 2. Relationships between quadrat height and width and the laser beam angle θ.
Calculating bottom slope and transect length
We will record data on the water depth at different points along the transect. Our horizontal
position will be determined by the ship's GPS. We will then calculate our position along the
transect using the following formula:
D  (lat 2  lat1 ) 2  ((lon 2  lon1 ) cos(lat avg )) 2 .
D is the distance in nautical miles between two points along the transect. Lon is the longitude
(expressed in minutes), lat is the latitude, and latavg is average latitude. The cosine accounts for
the shape of the earth which causes lines of longitude to bend toward each other approaching the
poles. The bottom slope (depth (m)/horizontal distance (m)) is determined by plotting the
measured water depth versus the distance along the transect, after converting both measurements
to the same units, such as meters. (One nautical mile = 1852m. One foot = 0.3048m.) Calculate
the slope of this line by linear regression. (The "Add Trendline" function in Excel.) The transect
length can then be calculated by dividing the change in depth (measured with the pressure sensor
on the ROV) by the slope.
Analyzing the video data – week two
During the second week we will analyze the video data. We’ll calculate the width and length of
each transect. And, we’ll count and identify all of the rockfish we record on each transect.
We’ll also characterize the bottom type of each transect (e.g., sand, boulders, gravel).
ECSI 322 – Oceanography Laboratory - Manual
14
Draft report
For your draft report, you’ll calculate bottom slope from the GPS and water depth data. First,
use the latitude and longitude from the GPS data to calculate the distance between each location
where we took a water depth reading using the depth sounder. Then, plot the distance among
locations versus water depth. Calculate the bottom slope of each transect (in degrees).
Final report
Your task is to address the following questions. What is the most important type of habitat for
rockfish? What type of habitats should be protected as part of efforts to promote rockfish
recovery? Determine the relationship between bottom slope and rockfish density (rockfish m-2).
Assess how other aspects of bottom habitat might influence rockfish distribution. For example,
did you observe spatial relationships between fish and particular features of the bottom habitat?
Also determine the depth distribution of each rockfish species. Include a few references from the
literature on rockfish and marine protected areas to bolster your report.
References
Auster, P,J., R. J. Malatesta and C. L. S. Donaldson. 1997. Distributional responses to smallscale habitat variability by early juvenile silver hake, Merluccius bilinearis.
Environmental Biology of Fishes 50: 195–200.
Palsson, W. et al. 2009. The biology and assessment of rockfishes in Puget Sound. WA Dept of
Fish and Wildlife Report No. FPT 09-04. 208 pp.
ECSI 322 – Oceanography Laboratory - Manual
15
Pre-laboratory Report 2, 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
questions. You may write your answers in the space available and turn in this sheet.
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:___________
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. Water depth
is 90 m and the depth of the pycnocline is 5 m. What is the rate of respiration (dissolved oxygen
removal) in deep water of Bellingham Bay?
Respiration rate (μmol O2/L/day)_______________
Show your calculations here:
ECSI 322 – Oceanography Laboratory - Manual
16
ESCI 322 Lab Report 2 Part 1, Vertical profiles in Bellingham Bay
The Nooksack River enters Bellingham Bay at its northern end. It delivers approximately one
billion cubic meters of freshwater to Bellingham Bay each year. This significantly reduces the
salinity of the bay and affects the mean circulation in the bay as well. The Nooksack also
delivers nutrients and other dissolved solutes and organic matter. Thus, it strongly affects the
biology and chemistry of Bellingham Bay. Recently, regions of low dissolved oxygen (DO)
concentrations have been observed in the center of the bay during late summer. A similar pattern
of low oxygen (termed hypoxia) has been observed in other small embayments throughout Puget
Sound. Furthermore, there are plans to double the size and output of the Post Point Wastewater
Treatment Plant, which empties into Bellingham Bay, by 2014. How will this change in nutrient
and freshwater input to Bellingham Bay affect its nutrient and oxygen concentrations?
The objective of today's lab is 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 periods. 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:
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
ECSI 322 – Oceanography Laboratory - Manual
17
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.
Measuring phytoplankton pigments: The CTD has a fluorometer that measures in-situ seawater
fluorescence. This measurement is related to the concentration of chlorophyll in the water. But,
the in-situ fluorometer must be calibrated with laboratory measurements of chlorophyll from
discrete water samples. To perform this calibration we will measure the concentration of
chlorophyll on some of the samples we collect with the rosette. We will first extract the
pigments using acetone. Then, we will use a different type of fluorometer to measure the
concentration of the extracted algal pigments.
Survey methods: We will measure depth profiles of temperature, salinity, light, chlorophyll and
dissolved oxygen at several locations in Bellingham Bay using a CTD. The CTD (conductivity,
temperature, depth) is the oceanographer’s primary sampling device. It consists of a set of
electronic probes attached to a metal rosette wheel 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).
Dissolved oxygen: The dissolved oxygen content of water is influenced mainly by water
temperature (cold water can hold more dissolved gas than warm water) and biological activity.
Primary producers (including macroalgae and phytoplankton) add oxygen to the water as they
photosynthesize. Recall that photosynthesis can only take place at depths shallow enough to
receive light. Aerobic organisms (plants, animals, aerobic microbes) consume oxygen during
metabolism. Waters containing high levels of organic matter (i.e. dead cells, organic-rich muds,
dissolved organic matter from sewage or other sources) may have low levels of oxygen because
heterotrophs use up the oxygen while decomposing the organic matter.
Water samples: Use a clean bucket firmly affixed to a line to obtain a surface water sample from
the shallow stations. Use a clean bucket to collect surface water samples at shallow stations. At
deeper stations, collect water from the CTD rosette. Close (“fire’) the bottles electronically
using the CTD software at selected depths. To sample the Niskin bottles, turn the knob at the top
ECSI 322 – Oceanography Laboratory - Manual
18
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.
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 dissolved oxygen
lowest?
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:
TO 
Si
R
Si  SO
ECSI 322 – Oceanography Laboratory - Manual
19
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:
Ti  T0  R
And, the bottom-water residence time (RT) can be calculated as follows:
RT 
VB
Ti
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).
Draft report: Plot the locations of each station in Bellingham Bay using any program you like.
Plot profiles of temperature, salinity, dissolved oxygen and density (t, calculated using formula
in the provided spreadsheet. Also, create a contour plot of surface salinity.
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.
ECSI 322 – Oceanography Laboratory - Manual
20
Figure 1. Station locations for Bellingham Bay survey
Figure 2. Predicted tidal levels in Bellingham Bay, October 18, 2011.
ECSI 322 – Oceanography Laboratory - Manual
21
Figure 3. Additional chart of Bellingham Bay
ECSI 322 – Oceanography Laboratory - Manual
22
Pre-laboratory Report 3: Seawater fluorescence, chlorophyll, and
phytoplankton biomass
Name: _______________________________
Read the description of this week’s laboratory assignment and access the Excel spreadsheet
sed_profiles.xls found on the course web site. Then, answer the following questions to be turned
in at the beginning of the lab period. You do not need to type your answers to these questions.
You may write your answers in the space available and turn in this sheet.
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
Surface sample
Deep sample
Blank reading
First fluorescence reading
17000
18000
208
Second fluorescence reading
10000
16000
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?
ECSI 322 – Oceanography Laboratory - Manual
23
ESCI 322 Lab Report 2, Part 2, Seawater fluorescence, light, chlorophyll, and
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).
ECSI 322 – Oceanography Laboratory - Manual
24
1) Methods for measuring irradiance and seawater fluorescence
Secchi disk:
A black and white disk is a low-tech way to measure the penetration of light into water. Named
for Father Pietro Angelo Secchi, this primitive instrument has been used extensively in marine
and aquatic systems for studying irradiance. Today, when the quantum sensor smashes on the
side of the ship, the Secchi disk comes out. Lower the disk until it can no longer be seen, then
raise it to the depth at which it first becomes visible. Record this depth. You may have to repeat
this several times to obtain a consistent Secchi depth.
Spherical light sensor (4pi):
This spherical light sensor measures the total amount of photosynthetically active radiation
(PAR, radiation at wavelengths that can be used in photosynthesis). PAR wavelengths range
from 400 to 700 nm for most photosynthetic organisms. Although light enters the water from
above, it is scattered by water molecules so that, from an underwater vantage point, light comes
from all directions. The spherical (4pi) sensor allows accurate measurement of this diffuse light
field. Irradiance is reported in units of µmoles photons m-2s-1. Obtain an irradiance profile by
lowering the sensor (carefully!) to discrete depths as indicated by markings on the cable. (Note
that the same data logger can be used with a 2 pi [flat, circular] sensor to measure the amount of
light striking the water’s surface, either instantaneously, or integrated over time.)
In situ fluorometer:
Chlorophylls absorb light energy at one wavelength and emit it a longer wavelength; this
property is known as fluorescence. The fluorometer works by shining blue light onto a sample
(called excitation) and measuring the resulting emission of red light. Filters are used to control
the wavelengths received by the sample and the detector (a photomultiplier tube). An in situ
fluorometer measures the fluorescence of whole seawater. By picking the appropriate excitation
and emission wavelengths, the measured seawater fluorescence should be related to the amount
of chlorophyll in the water. However, other water properties also fluoresce. Therefore, the
fluorescence data must be calibrated by comparing them to measured chlorophyll concentration
in discrete samples.
2) Collection of water samples for chlorophyll and (later) nutrient analysis.
Determine the depth of the pycnocline and the depth of maximum fluorescence using data
collected electronically by the CTD. Then, collect water samples from six depths using the
water sampling bottles on the CTD. Collect samples from above and below the pycnocline and
at depths with different fluorescence levels. Once on deck, remove water from the CTD rosette
bottles and fill one plastic sample bottle with water from each bottle; label with depth and station
ID. Filter the samples on deck, store in aluminum foil, and place into a cooler. These samples
will be frozen for later analysis in the laboratory.
3) Calculation of extinction coefficients
From Secchi disk data:
Light in water is absorbed and scattered by the water molecules themselves, and by dissolved
and particulate material in the water. The amount of light absorbed per unit surface light per
ECSI 322 – Oceanography Laboratory - Manual
25
meter depth is given by the extinction (or attenuation) coefficient k (m-1). The light attenuation
coefficient, KD, is calculated from the Secchi depth (ZSD) according to the formula KD=1.44 /ZSD.
From light meter data:
The extinction coefficient KD can also be calculated from vertical profiles of irradiance such as
those you obtained using the light meter. In this case, light (I) is assumed to decrease
exponentially with depth (z) according to the formula IZ = I0 e-kz. Based on this model, the
extinction coefficient can be calculated from a near-surface (I0) and a deep (IZ) irradiance
measurement and the depth interval (z) between these two measurements:
k = -1/z ln(IZ/I0). Note that the shallowest irradiance measurements are often ‘noisy’, so you
probably will not want to use these for this two-point method of calculating k. The most
accurate way to estimate k is to use all your data from a given vertical profile. Create a plot of ln
irradiance vs. depth (analogous to a semi-log plot for estimating growth rate). The slope of the
regression of ln irradiance vs. depth is -k.
4) Light absorption by algal pigments
All photosynthetic organisms contain pigments (Fig. 2) to harvest light energy and to protect
themselves from light-induced damage. Spectrophotometry takes advantage of light absorption
by pigments to estimate their concentration in a given sample. Within a certain range of
concentration, the absorbance of light is proportional to the concentration of pigment present
(Beer’s Law). The spectrophotometer passes a beam of light through a substance (in our case, an
organic extract) and the amount of light absorbed from the beam by the sample is determined.
The photo-diode array spectrophotometer is able to quantify absorbance over a range of
wavelengths simultaneously. We will examine absorption spectra from several different kinds of
pigments using the diode array spectrophotometer, including extracts of the pigments from our
study site. This kind of spectrophotometry can used quantitatively to determine pigment
concentrations in a water sample (see Parsons et al 1984 and Jeffrey et al. 1997 for
methodologies).
Fig. 2. Diagram of a cryptophyte cell. These are common members of marine phytoplankton
communities. The large chloroplast is bounded by membranes (CE, CER) and filled with layered
thylakoid membranes (T). Most of the pigments are imbedded in the thylakoid membranes (Lee
1999).
ECSI 322 – Oceanography Laboratory - Manual
26
Fig. 3. Absorbance spectra of some commonly occurring chlorophylls and carotenoids.
5) Measurement of pigment concentrations by fluorometry
Chlorophyll a fluorescence can be used to quantify the amount of chlorophyll present in the
particles in a water sample. This method is very sensitive, so it works well for dilute systems
such as the ocean. The fluorometer works by shining blue light onto a pigment extract and
measuring the resulting emission of red light. Filters are used to control the wavelengths received
by the sample and the detector (a photomultiplier tube). The amount of blue light used to excite
the fluorescence will influence the amount of fluorescence produced; this is controlled by a
series of “doors” and must be accounted for in the calculations. The fluorometer is standardized
using pure chlorophyll a extracts which in turn are quantified on the spectrophotometer (this has
been done for you). There are four steps involved in the measurement of water column
chlorophyll concentrations: i) filtering the water sample; ii) 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
ECSI 322 – Oceanography Laboratory - Manual
27
(this will be saved for nutrient analysis).
d) Label the nutrient sample bottle with the sample name.
e) Place the filter in the appropriate 15-ml centrifuge tube. This filter now has on it all the
particles (including chlorophyll-containing phytoplankton cells) that were in the original 50-ml
water sample.
Step 2: Sonicating the filter
a) Place a filter from step i into a 15-ml centrifuge tube and add 5 ml cold 90% acetone.
b) Sonicate while submerging the centrifuge tube in an ice-water bath for one minute. Wear
gloves, safety goggles and ear protection.
c) When the sample is thoroughly sonicated, add acetone until the final volume is 10 ml.
d) Record the final volume of solvent plus homogenate in the tube. This is your “extraction
volume”. Put the tube into a test tube rack for storage in the ice bath (or freezer for longer-term
storage).
Step 3: Measuring the sample fluorescence
a) Vortex or vigorously shake each centrifuge tube, then remove the filter, squeezing out any
solvent using a clean (solvent-rinsed) pair of forceps. Centrifuge the tubes (high speed, 5 min).
b) If extracts are visibly green, they must be diluted or the detector response will be saturated.
Use calibrated centrifuge tubes and automatic pipettes to dilute samples with 90% acetone; keep
track of all dilutions.
c) Zero the fluorometer using a cuvette containing 90% acetone. Re-zero every time you switch
door (sensitivity) settings.
d) Transfer your extract to a clean glass cuvette. Be careful not to resuspend any of the palletized
filter debris (this will interfere with the fluorescence reading).
e) Read the fluorescence of your extract. This value should be >25 and <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) =
K x Fm v( F0  Fa )d
v f ( Fm  1)
Phaeopigments (µg/liter seawater) =
K x Fm v( Fm Fa  F0 )d
v f ( Fm  1)
where:
Fo = fluorescence before acidification
ECSI 322 – Oceanography Laboratory - Manual
28
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.
ECSI 322 – Oceanography Laboratory - Manual
29
Pre-laboratory Report 4, 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
questions. You may write your answers in the space available and turn in this sheet.
1: What kinds of phytoplankton depend upon silicate as a nutrient?
2. You are measuring 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
0 µM
10 µM
50 µM
100 µM
200 µM
Absorbance
3.1
7.4
23.1
44.2
86.0
Absorbance reading of sample with unknown concentration: 76.1
ECSI 322 – Oceanography Laboratory - Manual
30
ESCI 322 Lab Report 3: Nutrients in Seawater
The objective of this laboratory session is to learn standard techniques for the analysis of
dissolved phosphate and nitrate. These nutrients often limit the productivity of coastal marine
systems. We will obtain data on silicate, phosphate, nitrate, and ammonia 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
ammonia, 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 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.)
ECSI 322 – Oceanography Laboratory - Manual
31
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. Add 1 ml
chloroform, store refrigerated. Stable for many months.
Secondary standards:
2° STD (M)
0.00
0.75
1.50
2.25
3.00
5.50
L 1° STD/50 ml H2O
0
25
50
75
100
150
Quality control standard:
1.50

Procedure:
1. Pipette 5 ml aliquots of D.I. water blanks (3x), analytical and quality control standards (3x), 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 filings coated with metallic copper.
The nitrite produced is determined by diazotizing with sulfanilamide and coupling with N-(1-naphthyl)ethylenediamine to form a highly colored azo dye which can be measured spectrophotometrically. Any
nitrite initially present in the sample must be corrected for.
Nitrate standards
Primary standards – 10mM potassium nitrate solution. Dissolve 0.76 g potassium nitrate in 250
ml Nanopure water plus two drops of chloroform. Refrigerate.
ECSI 322 – Oceanography Laboratory - Manual
32
Reagents have also been made ahead of time. This procedure uses five: 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.
Draft report
Calculate the concentrations of ammonium and nitrate (+ nitrite) in the samples we collected
from Bellingham Bay and Burrows Bay. Plot these data as profiles.
Part II, the following week: Phosphate and silicate
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
using 10-cm cuvettes.
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 50M
SIO4). Ordinarily, these would be made by dissolving Na2SiF6 in DIW.
ECSI 322 – Oceanography Laboratory - Manual
33
Reagents: Reagents have also been made ahead of time. This procedure uses three: Metolsulfite 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 [0M 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
Metol-sulfite
Oxalic Acid
H2SO4
DIW
1 sample
0.2 ml
0.12 ml
0.12 ml
0.16 ml
10 samples
2 ml
1.2 ml
1.2 ml
1.6 ml
20 samples
4 ml
2.4 ml
2.4 ml
3.2 ml
50 samples
10 ml
6 ml
6 ml
8 ml
100 samples
20 ml
12 ml
12 ml
16 ml
7. Add 1.5 ml of the mixed reducing reagent to all samples and standards. Cap tightly and
shake once or-twice. Wait at least 2.5 hr for the yellow silicomolybdic acid to be
reduced to a deep blue "silicomolybdous acid complex" - whose exact formula isn't
known. In samples whose [Si(OH)4] is < ~1.5 M it will be difficult to see any color, but
all others should be visibly blue. (If samples are not visibly blue, the waiting period can
be reduced to 1.5 hr.)
While you are waiting for this reaction to occur, proceed to the phosphate measurements.
8. Now, add 1.0 ml of acidified ammonium molybdate reagent to the blank. This procedure
is called a reverse-addition blank. The oxalic acid in the mixed reducing agent blocks the
formation of the yellow silicomolybdic acid so none forms when the acidified ammonium
molybdate reagent is added later.
9. Measure the absorbance at a wavelength of 810 nm in a spectrophotometer (the blue
complex absorbs most strongly in the infrared), using glass culture tubes. We can use
glass at this point because the silicate that we will measure has already formed the blue
silicomolybdous acid complex and the oxalic acid blocks further complex formation.
ECSI 322 – Oceanography Laboratory - Manual
34
Calculations:
The [Si(OH)4] of each sample is calculated as:
[Si(OH)4] (M)
= Sample absorbance - Reverse-addition blank absorbance
Slope of standard curve (absorbance/M)
I will provide you with an excel spreadsheet that will help you make the appropriate calculations.
Dissolved phosphate (H2PO4-, HPO42-, PO43-) determination (Parsons et al. 1984)
Phosphate is an important limiting nutrient in freshwater and marine systems. Eutrophication in
freshwater systems are often the result of high inputs of P, which lead to increased growth of
algae, and sometimes to reductions in dissolved oxygen or other problems. The importance of
phosphate for regulating the growth of algae was spectacularly demonstrated by Vollenweider
(1976). Figure 1 shows the relationship between total P loading and algal biomass in lakes.
Figure1. Empirical model relating mean surface chlorophyll-a concentrations of lakes to phosphorus
loading, adjusted for hydraulic loading. Plot from Cloern, 2001. Data from Vollenweider, 1976.
Outline of method:
The seawater sample is allowed to react with a reagent containing molybdic acid, ascorbic acid,
and trivalent antimony. The resulting complex is reduced to give a blue solution that is
measured at 885 nm.
Procedures:
Samples should be at room temperature. Turn on Spectrophotometer and set wavelength to 885
nm. This is the light absorption maximum for the blue solution produced in this method.
Standards: To save time we will make a set of working standards from a secondary standard that
has been prepared in advance. First a primary standard solution was made from anhydrous
KH2PO4 and DIW. The secondary standard was made from the primary standard.
ECSI 322 – Oceanography Laboratory - Manual
35
Label and rinse five 100-ml volumetric flasks several times with DIW. Fill them with about 90
ml of DIW. Then, rinse an Erlenmeyer flask several times with a few ml of the secondary
standard solution. Using a 5-ml pipette, carefully add the secondary standard solution from the
Erlenmeyer flask to each 100-ml volumetric flask according to the table below. Bring up the
volume to 100 ml by carefully adding DIW. These are your tertiary standards.
3° STD (µM)
0.0
0.6
1.8
3.0
4.2
ml 2° STD / 100 ml H2O
0.00
0.50
1.50
2.50
3.50
Place ten glass culture tubes in a tube rack (two for each standard) and record their positions in
your notebook. Also add five tubes for the seawater sub samples and record their positions.
Mixed reagent: We will create the mixed reagent from single reagents that were made in
advance of today’s lab.
For each 5-ml sample, use 2 ml Ammonium molybdate, 5 ml Sulfuric acid, 2 ml Ascorbic acid, and 1 ml
K-antimonyl-tartrate. First, wash a graduated cylinder with a few ml of ammonium molybdate three
times. Then, measure the correct volume of ammonium molybdate and the other reagents according to
the table below. Add these reagents to a labeled Erlenmeyer flask, cover with parafilm, and mix for 30
seconds. Use volumes in the 50 samples column.
Reagent
Ammonium molybdate
Sulfuric acid
Ascorbic acid
K-antimonyl tartrate
10 samples
2.0 ml
5.0 ml
2.0 ml
1.0 ml
20 samples
4.0 ml
10.0 ml
4.0 ml
2.0 ml
50 samples
10.0 ml
25.0 ml
10.0 ml
5.0 ml
Measure 5 ml of each sample, each STD, and the nanopure H2O blank into culture tubes.
Add 0.5 ml mixed reagent. Wait 5 minutes. Measure absorbance on a spectrophotometer.
Spectrophotometric Procedures
Set the spectrophotometer at 885 nm. Turn spec on and allow to warm up for 15 min. Set
wavelength to 885 nm. Set filter lever to the right. Set to 0% transmittance. Change mode
to “absorbance”. Insert Nanopure water blank (this is not the same as the reagent blank). Set
to 0% absorbance. Insert sample and record absorbance.
Data Analysis for Phosphate
Follow the data analysis procedures for silicate.
Reporting nutrient concentrations in Bellingham Bay
The spatial distribution of temperature, salinity, and nutrients changes every time I sample Bellingham
Bay with students from this class. This is because each time we sample it at different stages of the tide
ECSI 322 – Oceanography Laboratory - Manual
36
and under different wind conditions, and because the Nooksack River flow varies from year to year.
One way to make sense of the nutrient concentrations we measure, however, is to plot them versus
salinity. This enables us to examine the relationships between river input from the Nooksack, salt water
input from the Strait of Georgia and the Strait of Juan de Fuca, and nutrient concentrations without
reference to specific locations within the bay. We can even infer whether biological or chemical
processes produce or consume nutrients within Bellingham Bay as well. This method will be described
next.
Understanding relationships between salinity and nutrient concentrations
Tracer concentration
A
Tracer concentration
Mixing theory: A non-reactive solute (sometimes referred to as a conservative tracer) input from a river
will be diluted by seawater. If dilution is the only process that determines the concentration on such a
tracer in an estuary, the relationship between tracer concentration and salinity will be linear. This also
holds for a tracer with a seawater input.
River source
B
Marine source
Tracer concentration
Salinity
Salinity
A
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.
B
Estuarine source
Salinity
Figure 1. Theoretical
relationship between a
conservative solute and
salinity for solutes with
(A) a river source and
(B) a marine source.
Estuarine sink
Salinity
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.
ECSI 322 – Oceanography Laboratory - Manual
37
DIN:DIP ratio
N > P:
Nitrogen fixation?
16
8
N > P:
Phosphate storage
In sediments?
Redfield ratio
N < P:
Fertilizer input?
0
Salinity
N < P:
Denitrification
In sediments?
Figure 3. Several processes that can cause nutrient
concentrations to deviate from Redfield proportions. (This list
is not complete.) Deviations at low salinity may reflect river
inputs. For example, phosphate-based fertilizers reduce the
N:P ratio whereas freshwater N fixation can increase it.
Deviations at high salinity in an estuary may reflect processes
affecting bottom waters. For example, denitrification in marine
sediments can reduce N:P whereas sorption of P onto
sediments could potentially increase the ratio. Also,
preferential uptake of N versus P in the marine waters entering
Bellingham Bay could reduce the N:P ratio.
Creating a nutrient budget for Bellingham Bay: The data we have collected will allow us to
calculate a nutrient budget. This is a careful accounting of the rates of supply of nutrients from different
sources. The four potentially important sources that we can compare are: the Nooksack River, the Post
Point WWTP, nutrient supply from sediment, and deep water inflow from the Strait of Georgia/Strait of
Juan de Fuca. For the sediment component of the budget, divide the nutrient flux by the water depth as
we did for the sediment oxygen consumption rate. For the other components, we need to multiply the
nutrient concentrations in water from the Nooksack River, Post Point WWTP, and deep water with their
volumetric flow rates. Example: (15 mmole/m3 Nitrogen) * (7,000,000 m3/day Nooksack R. water) =
1.05*108 mmole/day nitrogen from the Nooksack River.
Create a budget for both nitrogen and phosphorus. You’ll need to calculate total nitrogen as the sum of
ammonium, nitrate and nitrite. Flow rates for the Nooksack River are available from the USGS. Flow
rates for the Post Point WWTP are available too. We can calculate the volumetric flow rate of deep
water from the Nooksack River flow rate and the extent to which Bellingham bay water is diluted 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 and compare your
data with theoretical expectations. Present your nutrient budget as a table in your report.
References
Cloern, J. E. 2004. Our evolving conceptual model of the coastal eutrophication problem. Mar. Ecol.
Prog. Ser. 210, 223-253.
Parsons, T. R., Y. Maita, and C. M. Lalli. 1984. A manual of chemical and biological methods for
seawater analysis. Pergamon Press, Elmsford, N.Y.
ECSI 322 – Oceanography Laboratory - Manual
38
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.
ECSI 322 – Oceanography Laboratory - Manual
39
Pre-laboratory Report 5, Phytoplankton growth and grazing
Name: _______________________________
Read the description of this week’s laboratory assignment and answer the following questions to
be turned in at the beginning of the lab period. You do not need to type your answers to these
questions. You may write your answers in the space available and turn in this sheet. But, you’ll
also need to turn in a data plot and linear regression described below.
Problem:
You conduct a dilution experiment. After making the dilution series shown below, you allow
phytoplankton to grow for 24h and then stop the experiment by filtering the phytoplankton. You
measure the concentration of chlorophyll-a in the filtered samples. The data you collect are
shown in the table below.
Table. Hypothetical results from a dilution experiment
Dilution factor
(fraction seawater)
1.00
0.75
0.50
0.25
0.05
Initial chlorophyll
concentration (p0)
30
22
11
3.0
0.15
Final chlorophyll
concentration (p)
38.5
53.3
47.1
29
4.0
Net growth
1/t * ln(p/p0)
1. Fill in the last column of the table [ 1/t ln(p/p0) ].
2. Calculate the slope and intercept from the linear regression of the net growth versus dilution
factor. (Show the Excel plot you created including the data points and linear regression line.)
Slope: _____________
Intercept: __________
3. What are the per-capita growth and grazing rates that you determined from this experiment?
Growth rate: ________________
Grazing rate: ________________
ECSI 322 – Oceanography Laboratory - Manual
40
ESCI 322 Lab Report 4: Phytoplankton population growth and grazing
Part 1: Setting up the experiment
The abundance of phytoplankton in the ocean is set by the balance between rates of population
growth, mortality and transport. Phytoplankton population growth rates in the field are affected
by light, nutrient concentration, temperature, and species composition among other factors.
Consumption by grazers is one of the largest sources of mortality. There are many grazers of
phytoplankton in Puget Sound; herring, calanoid copepods and bottom-dwelling filter feeders
such as clams are relatively large grazers. There are also phytoplankton grazers that are barely
larger than the phytoplankton cells they consume. These protistan grazers are termed
microzooplankton. In many marine ecosystems, microzooplankton are extremely important
phytoplankton consumers. The objective of this lab is to measure rates of phytoplankton growth
and microzooplankton grazing. We’ll perform the experiment at two light levels to better
understand how light and grazing interact to control phytoplankton abundance in the sea.
Theory: Consider the equation for population growth. It can be written in differential form as
dp
follows:
 p , where p is the concentration of phytoplankton and µ is the per-capita growth
dt
rate. If z is the microzooplankton concentration and χ is the phytoplankton-zooplankton
dp
encounter rate, you can add grazing to this equation as follows:
 p  zp . Now, consider
dt
what would happen if you diluted a sample of seawater with filtered seawater. Dilution should
not affect the growth rate of phytoplankton which multiply by binary fission. However, dilution
will reduce the encounter rate between phytoplankton and zooplankton, lowering the grazing
rate. If you add dilution (D is the fraction of undiluted seawater in the sample.) to the population
dp
growth model, you can write it as
 p  gDp , where g is the per capita mortality rate of
dt
phytoplankton due to grazing. The solution to this model is p = p0e(µ-gD)t, where p0 is the initial
phytoplankton concentration. Taking the natural log of both sides and rearranging this equation
1  p 
    gD . The first term is a measure of phytoplankton
gives the following formula: ln 
t  p 0 
net growth rate. Plotting this versus the dilution factor, D, yields a straight line with a slope
equal to g and intercept µ (Fig. 1). Thus, if you conduct a dilution experiment you can calculate
both the phytoplankton per capita growth and grazing rates. In our experiment, we will conduct
a dilution experiment with two light intensity treatments (full-strength light and 50% light).
We’ll use mesh screening material to manipulate light intensity and measure phytoplankton
growth and grazing rates at the two light levels. This will allow us to ask the question how does
phytoplankton growth respond to light and grazing?
Methods: It is a challenge to work with phytoplankton in late fall when biomass approaches its
lowest levels due to low light levels. So, I will collect a big sample of seawater and let the algae
grow under well-lit conditions in the laboratory for a week prior to our class. That should give
us a sample with a high enough biomass to use for our experiment. Filter the water to create
three dilution levels – full-strength seawater, 50% seawater diluted with filtered seawater, and
10% seawater diluted with filtered seawater. Perhaps the trickiest part of setting up this
ECSI 322 – Oceanography Laboratory - Manual
41
experiment will be filtering the water. You’ll need to filter a large quantity of water without
damaging the phytoplankton cells and then divide the seawater into replicate bottles while
keeping the cells in suspension so that all bottles receive the same concentration of cells. Filter
half the seawater through a filter cartridge using a peristaltic pump which won’t damage the
plankton. While filtering, use a piston to keep the cells in suspension. Then, combine the whole
and filtered seawater to create the following treatments, keeping the cells in suspension while
doing so.
Fraction whole
Seawater
100%
100%
100%
50%
50%
10%
10%
10%
Number of
replicates
4
2
2
3
1
4
2
2
Light
level
NA
100%
10%
NA
100%
NA
100%
10%
Sampling
time
initial
final
final
initial
final
initial
final
final
Incubate the samples for 48 hours. End the experiment by preserving a 15-ml sample from each
bottle using Lugol’s iodine preservative and pass the rest of the samples through a glass fiber
filter. Freeze the filters for analysis next week.
Dilution experiments
Y-intercept= “infinite dilution”
Net Growth Rate (per time)
GROWTH RATE
Growth equation: dp/dt = p – zp
dp/dt = p – zDp = p – gDp
Solution: 1/t ln(p(t)/p0) =  – gD
Slope = linear relationship
with dilutions
HERBIVORY RATE
Decreasing # of Grazers
Dilution factor (fraction SW)
0
1
Figure 1. Theoretical relationship between net growth rate and dilution factor. The slope gives
the per capita grazing rate and the y intercept gives the per capita growth rate.
Reference: Landry, M.R., 2001. Microbial loops. In: Steele, J.H., Thorpe, S., Turekian, K.
(Eds.), Encyclopedia of Ocean Sciences, Academic Press, London, pp. 1763–1770.
ECSI 322 – Oceanography Laboratory - Manual
42
ESCI 322 Lab Report 4: Phytoplankton population growth and grazing
Week 2: Sample processing and analysis
Reconsider the population growth equation that forms the basis of our dilution experiment:
1  p 
    gD . Note that ln(p/p0) is unitless. It depends on the ratio of p/p0 (the ratio of
ln 
t  p 0 
final to initial phytoplankton concentration) and the units cancel. This means we can use any
measure of concentration we like to quantify the phytoplankton. The units don’t matter. We will
use chlorophyll concentration since it can be measured relatively easily and precisely. We will
also enumerate phytoplankton and microzooplankton cells to determine which species were most
strongly affected by grazers.
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.
ECSI 322 – Oceanography Laboratory - Manual
43
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) =
K x Fm v( F0  Fa )d
v f ( Fm  1)
Phaeopigments (µg/liter seawater) =
K x Fm v( Fm Fa  F0 )d
v f ( Fm  1)
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
ECSI 322 – Oceanography Laboratory - Manual
44
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. This will give the number of cells per 10 ml (because of the 10-fold concentration
step).
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.
ECSI 322 – Oceanography Laboratory - Manual
45