Aquatic Field and Lab II

Aquatic Field and Lab II
NCSR
EDUCATION FOR A SUSTAINABLE FUTURE
www.ncsr.org
AQU
ATIC FIELD AND
QUA
LAB METHODS II
NORTHWEST CENTER FOR SUSTAINABLE RESOURCES (NCSR)
CHEMEKETA COMMUNITY COLLEGE, SALEM, OREGON
DUE # 0101498
FUNDING PROVIDED BY THE NATIONAL SCIENCE FOUNDATION
OPINIONS EXPRESSED ARE THOSE OF THE AUTHORS AND NOT
NECESSARILY THOSE OF THE FOUNDATION
Aquatic Field and Lab Methods II
The Northwest Center for Sustainable Resources is an Advanced
Technological Education project funded by the National Science
Foundation.
Aquatic Field and Lab Methods II was developed at Grays Harbor College, Aberdeen, Washington, and was tested and revised at Mount Hood
Community College, Gresham, Oregon. Materials were prepared by
Don Samuelson, Lead Program Developer for NCSR. Samuelson holds
a M.S. in General Science/Fisheries from Oregon State University, a
B.A. in Chemistry/Biology from Pacific Lutheran University, and an
A.S. in Marine Biology from Grays Harbor College.
Technology education programs in which this course is incorporated
are described fully in the Center’s report entitled, “Visions for Natural
Resource Education and Ecosystem Science for the 21st Century.”
Copies are available free of charge.
The authors and the Center grant permission for the unrestricted use
of these materials for educational purposes. Use them freely!
Course materials are also posted on our website:
www.ncsr.org
Please feel free to comment or provide input.
Wynn W. Cudmore, Ph.D., Principal Investigator
Northwest Center for Sustainable Resources
Chemeketa Community College
P. O. Box 14007
Salem, OR 97309
Phone: 503-399-6514
E-mail: [email protected]
Table of Contents
FISH 221
COURSE OUTLINE ................................................................................................................... 1
LABS & ACTIVITIES
FIELD NOTEBOOKS
AND
JOURNALS .......................................................................................... 1 1
PLANKTON SAMPLING AND ANALYSIS ........................................................................................ 1 5
NOTES FOR INSTRUCTORS ....................................................................................................... 2 2
BIOMETRICS ............................................................................................................................ 2 5
STREAM ANALYSIS—ELECTROSHOCKING ................................................................................... 3 6
NOTES FOR INSTRUCTORS ....................................................................................................... 4 2
STREAM ANALYSIS—DISCHARGE .............................................................................................. 4 5
MACROINVERTEBRATE SAMPLING .............................................................................................. 5 0
NOTES FOR INSTRUCTORS ....................................................................................................... 5 6
MARK-RECAPTURE METHOD .................................................................................................... 5 8
GAS SUPERSATURATION ........................................................................................................... 6 5
NOTES FOR INSTRUCTORS ....................................................................................................... 6 8
BEACH SEINING AND OTTER TRAWLING ................................................................................... 7 4
NOTES FOR INSTRUCTORS ....................................................................................................... 8 2
NEKTON SAMPLING ................................................................................................................. 8 4
CAPSTONES & SEMINARS
CAPSTONE I: TEAM BUILDING .................................................................................................. 8 8
NOTES FOR INSTRUCTORS ....................................................................................................... 8 9
CAPSTONE II: ECOSYSTEM STUDY ............................................................................................ 9 0
NOTES FOR INSTRUCTORS ....................................................................................................... 9 1
WEEKLY SEMINAR ASSIGNMENT #1 ...................................................................................... 103
WEEKLY SEMINAR ASSIGNMENT #2 ...................................................................................... 104
STUDENT ASSESSMENTS
QUIZ I. ............................................................................................................................... 106
QUIZ II ............................................................................................................................... 107
SAMPLE MID-TERM EXAM .................................................................................................... 109
ORAL EXAM
AND
STUDY GUIDE ........................................................................................... 111
SAMPLE FINAL EXAM ........................................................................................................... 114
FISH 221
Course Outline
Course Outline
INTRODUCTION
This course has been adapted for use by the Northwest Center for Sustainable Resources. It is
suitable for undergraduate and community college programs as well as secondary school programs.
Aquatic Field and Lab Methods II was designed to be a “capstone/learning community” course for
final quarter sophomores majoring in natural resources and fisheries technology. As a follow-on to
Aquatic Field and Lab Methods I (a 6 credit capstone course taught the previous quarter), this
course continues to build on mastery of water quality data collection, analysis and presentation
skills. This course also addresses the learning of biological sampling, analysis and presentation.
The first five (5) weeks of the quarter, are spent learning biological sampling techniques (i.e.,
plankton analysis, electroshocking, macroinvertebrate sampling, beach seining and otter trawling).
Each student is assigned approximately 2-3 different methods so they can become “expert” in these
methods. Once they master these, they are responsible for teaching them to the other students in
the class. Mastery of the methods is then demonstrated to the instructor.
Also, during the first half of the quarter the class reads and seminars on a wide range of topics
including ecosystem management, understanding ecosystems, stream channels, large woody debris
in forested streams, stream temperature and aquatic habitats. The class begins learning and using
some project planning and management tools outlined in “The Memory Jogger II—A Pocket Guide
of Tools for Continuous Improvement and Effective Planning.” Some initial experimental design
work for the ecosystem study will be initiated in the last half of the course.
Between weeks 6-11 a comprehensive ecosystem study is carried out at a nearby field research
site. An overnight ROPES/Challenge course is the first activity to develop the self confidence,
self esteem, leadership and teamwork necessary to complete the ecosystem study. Students are
required to keep field notebooks and formal journals during the entire quarter. Project planning is
well underway by weeks 6-7. Project management continues from week 7-11. As the class takes
on the roles of their chosen study, mission, goals and objectives, scope-of-work, and task and
timelines are developed to complete the work plan.
The ecosystem study includes conducting a physical, chemical and biological baseline study at the
field site. Students begin weekly monitoring of such characteristics as plankton in a lake, and
macroinvertebrate populations in a creek. In addition, students use nets and electroshockers to
sample fish populations in the lake and creek. Water quality sampling (using techniques learned
during the previous quarter in Aquatic Field and Lab Methods I) could also be programmed into
the weekly sampling schedule.
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Course
Outline
Products of this capstone course include: keeping a field notebook and journal; completing several
problem sets (biometrics); participating in eight (8) seminars and facilitating at least one (1) of
them; completing a team-prepared written research report of ecosystem study results; writing an
article for a local newspaper; preparing a video of the learning community experience; attending
one (1) natural resource meeting in the community; making two (2) oral presentations to the
general public and a professional audience; and turning in a portfolio with work samples. In
addition, students are required to write self evaluations at the end of the quarter, and students will
evaluate their peers.
This is a 6-credit course which meets for 3 lecture hours and 6 lab hours per week. Aquatic Field
and Lab Methods II allows students to learn the basic concepts, techniques and skills used in
ecological surveys and research. The scientific method, experimental design, data collection,
recording, analysis and presentations are emphasized. Basic statistical analyses using computers
are routinely used in this course, and field training includes small boat handling and the skilled use
and care of biological sampling equipment. Students work individually and in teams to safely
carry out “real world” lake and stream surveys. Written research reports and students oral
presentations are major components of the course.
TEXT AND REFERENCES
DePree, M. 1992. Leadership Is An Art. Del Publishing, a division of Bantam Doubleday Del
Publishing Group, Inc. New York, NY. ISBN: 0-440-50324-8
Cox, G.W. 1996. Laboratory Manual of General Ecology. Seventh Edition. W.C. Brown Publish
ers. 278 pp.
Herman, S.G. 1994. The Naturalist’s Journal – The Grinnell System of Journel Writing. The
Evergreen State College. 275 pp.
COURSE DESCRIPTION
The primary goal of this field and laboratory course is for students to become skilled in the basic
use of biological techniques and equipment that are used at the entry-level of employment in
fisheries and natural resources. Along with learning the responsible, ethical, and safe use of these
field and lab instruments and tools, students will be able to maintain them, troubleshoot and make
minor repairs. Data collection, analysis and reporting of results (in both oral and written formats),
are major objectives of this course. Students will work both individually, and in teams, to
complete a wide array of learning outcomes.
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Course
Outline
COURSE OBJECTIVES
Upon successful completion of the course, students should be able to:
1. Demonstrate an understanding of the scientific method and experimental design in
actual field research studies.
2. Work individually and in teams to plan and execute a biological field survey. Project planning
will include mission, goals, objectives, scope of work, tasks and timelines.
3. Perform the following biological sampling methods with a high degree of knowledge, skill,
precision and accuracy:
a.
b.
c.
d.
e.
f.
plankton sampling.
beach seining.
otter trawling.
macroinvertebrate sampling.
mark-recapture population assessment.
various aspects of timber, fish and wildlife sampling and monitoring.
4. Work safely, both individually and in teams, in a supervised, “real world” lab and field setting.
5. Demonstrate critical thinking skills through active participation in seminars, project planning,
report writing and oral presentations of their research project.
6. Use previously learned problem solving, statistics and computer skills to evaluate and present
results of field and lab exercises. Critically think about how economic, political, and social
factors affect ecological policies and decision-making.
7. Maintain accurate, insightful, and timely field notebooks and journals throughout the term.
8. Creatively express themselves, orally and in writing, through field notebooks, journals,
seminars, and written and oral presentations of formal research results; be able to use a variety
of contemporary audio-visual techniques in their presentations.
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Course
Outline
STUDENT ASSESSMENT
Students may be assessed on a variety of activities which they have undertaken during the course.
Traditional assessment:
A.
B.
C.
D.
E.
F.
Quiz I – 25 pts
Quiz II – 50 pts
Mid-Term Exam – 100 pts
Oral Exam – 100 pts
Final Exam – 125 pts
Seminars – 16-20 pts each
Additional methods of assessment:
A.
C.
D.
E.
Field notebook and journals
Written research report
Presentation of field research to general and professional audiences
Demonstration of lab/field skills using the following equipment/methodologies:
1.
2.
3.
4.
5.
6.
Electroshocking
Surber Sampler
Ponar Dredge
Plankton Sampling/Counting
Beach Seining
Otter Trawling
F. Self-assessment of performance
G. Instructor evaluation of student performance (written)
H. Peer review of performance
TOPICS
I.
Introduction to Biological Field and Laboratory Methods
A.
Syllabus
1.
2.
3.
4.
5.
6.
7.
8.
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Course
Outline
Texts
Course Description
Course Objectives
Topics
Techniques of Instruction
Laboratory/Field Exercises
Methods of Evaluation
Other Instructional Materials, Equipment, Software and References
B.
Course Outline
1.
2.
3.
II.
Field Notebooks and Journals
Seminaring
Project Planning/Field Research Project
Overview of an ecosystem study
A.
B.
C.
D.
III.
Ecosystem management (seminar)
Literature research in ecology
Experimental design in ecological studies
Understanding ecosystems I (seminar)
Important considerations of ecological studies
A.
B.
C.
IV.
Sampling design in ecological studies
Writing an ecological research paper
Understanding ecosystems II (seminar)
Quantitative descriptions of ecological samples
A.
B.
C.
V.
Testing basic ecological hypothesis about samples
Stream channels (seminar)
Regression, correlation and analysis of variance
Management/leadership
A.
B.
C.
VI.
A.
B.
C.
D.
E.
F.
Personal time management
Continuous Quality Improvement (CQI)
Leadership Secrets of Atilla the Hun (seminar)
Using Memory Jogger II
“Changing Paradigms” (video)
Continuous Quality Improvement tools
Brainstorming
Developing an affinity diagram
Developing a tree diagram
Participating in the ROPES/Challenge course
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Course
Outline
VII.
A.
B.
C.
D.
Project planning (ecosystem study)
Defining our mission and scope-of-work
Using project planning software for ecosystem study tasks and time lines
How to conduct team meetings
Creating a cause and effect diagram
VIII.
Project management (ecosystem study)
A.
B.
C.
D.
E.
F.
G.
H.
IX.
A.
B.
C.
D
X.
Collection and analysis of physical, chemical and biological data
Process improvement (class activity)
Fine sediments
Large woody debris
Stream temperature and aquatic habitat
Creating a flow chart (class activity)
Nominal group technique (class activity)
Demonstration of ecological sampling skills (individual assessment)
Learning to work together
Conducting team meetings
Preparing for oral presentations
Editing and assembling final written report
Rehearsal for oral presentation
Assessment – Instructor’s choice of method from variety of assessment choices.
LABORATORIES AND ACTIVITIES
Field Notebooks and Journals
Plankton Sampling and Analysis
Biometrics—Terminology and Problems
Stream Analysis—Electroshocking
Stream Analysis—Discharge
Macroinvertebrate Sampling—Surber Sampler, Ponar Dredge, and Multiple-Plate Sampler
Mark-Recapture Method—Lincoln Index
Gas Supersaturation
Beach Seining and Otter Trawling
Nekton Sampling
Additional activities:
Capstone I—Team Building (The ROPES Challenge)
Capstone II—Ecosystem Study (Learning Communities)
Seminars: Special Topics and Internet Research
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Course
Outline
DETAILED SCHEDULE
Week 1
Lecture: Introduction to Biological Field and Laboratory Methods
Reading: Preface; Notes to Students; Chapter 1; pp 1-8 (lab text)
Lecture: Designing An Ecological Study; Ecosystem Management
Week 2
Lecture: Literature Research In Ecology
Reading: Chapter 3; pp 21-25 (lab text)
Lecture: Experimental Design in Ecological Studies
Reading: Chapter 4; pp 26-32 (lab text); Understanding Ecosystems
Seminar Topic: Understanding Ecosystems—Part I
Assignment: Biometrics problem
Week 3
Lecture: Sampling Design in Ecological Studies
Reading: Chapter 5; pp 33-39 (lab text)
Lecture: Writing An Ecological Research Paper
Reading: Chapter 6; pp 39-42 (lab text)
Seminar Topic:Understanding Ecosystems—Part II
Week 4
Lecture: Quantitative Description of Ecological Samples
Reading: Chapter 7; pp 43-51 (lab text)
Lecture: Testing Basic Ecological Hypothesis About Samples
Reading: Chapter 8; pp 53-59 (lab text)
Seminar Topic: Stream Channels—The Link Between Forests and Fishes
Week 5
Lecture: Regression, Correlation, and Analysis of Variance
Handout Topic: Time Management
Lecture: Personal Time Management
Seminar Topic: Fine Sediments and Salmon Production
Reading: Leadership Secrets of Atilla the Hun
Week 6
Reading: Continuous Quality Improvement (CQI) Tools; pp1-22
Class Activity: Brainstorming (Affinity Diagram)
Reading: Continuous Quality Improvement (CQI) Tools; pp 23-30; 156-160
Seminar Topic: Fine Sediments and Salmon Production
Reading: Memory Jogger II, pp 23-30; 56-62; 91-114; 150-155
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Course
Outline
Week 7
Class Activity: Defining Our Mission and Scope-of-Work
Reading: Memory Jogger II, pp 23-30; 154-155
Class Activity: Team Meeting #1; Cause and Effect (Fishbone)
Reading: Memory Jogger II, pp 56-62
Seminar Topic: Large Woody Debris in Forested Streams in the PNW: Past, Present, and Future
Week 8
Class Activity: Team Meeting #2; Flow Charts
Reading: Memory Jogger II, pp 115-131
Class Activity: Process Improvement Exercise
Reading: Memory Jogger II, pp 115-131 (review)
Seminar Topic: Stream Temperature and Aquatic Habitat—Fisheries and Forestry Interactions
Week 9
Class Activity: Team Meeting #3; Process Improvement
Assignment: Self Evaluations and Portfolios
Class Activity: Nominal Group Technique (NGT)
Assignment: Results of NGT
Seminar: Learning To Work Together
Week 10
Class Activity: Team Meeting #4
Class Activity: Final Preparation for Oral Presentation to General Audience
Class Activity: Rehearse for Oral Presentation to a Professional Audience
Class Activity: Team Meeting #5
Week 11
Finals Week: Instructor’s Choice of Assessment is conducted here
8
Course
Outline
INSTRUCTION TECHNIQUES
Lectures are supplemented with whiteboard illustrations, overhead projections, and seminars about
special reading assignments, discussions, and handout materials.
Laboratories include keeping an accurate field notebook and journal; use of active and passive fish
sampling devices (i.e., electroshocker; plankton, gill, and trawl nets; beach seines); collecting,
preserving, identifying and classifying biological specimens; and small boat handling; maintenance
and repair. Students work individually and in teams to carry out ecological research studies. They
are required to write and speak about their results.
OTHER MATERIALS, EQUIPMENT, SOFTWARE, AND REFERENCES
I. Sampling Equipment—Two (2) Boston Whalers, beach seines and otter trawls, electroshocker,
and a wide array of biological sampling equipment and supplies.
II. AV Equipment (available to students)
A. Overhead projector and mylar projections
B. 35 mm cameras and film or digital cameras
C. LCD projectors
D. Slide projectors
E. Video cameras and blank video tapes
III. Hardware, Software, Video
IBM PC’s (located in the classroom)
Word processing, presentation and spreadsheet software
Project planning software (PERT and Gantt)
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Course
Outline
REFERENCES
Robert, W. 1990. Leadership Secrets of Atilla The Hun. Warner Books. New York, NY.
Video, “Changing Paradigms,” from the book by Joel A. Baker, Discovering the Future: the
Business of Paradigms. 2nd Ed. Washington Society of Chartered Public Accountants. Registration
# 72514. Videotape distributed by Chart House Learning Corporation (1-800-328-3789).
Standard Methods for the Examination of Water and Wastewater. 1985. 17th Ed. American Public
Health Association (APHA), American Water Works Association (AWWA), and the Water
Pollution Control Federation (WPCF).
Statistix for Windows—User’s Manual. 1996. Analytical Software, P.O.B. 12185, Tallahassee, FL
32317-2185. 333pp.
Fisheries Techniques. 1995. Second Edition. American Fisheries Society. 732pp.
The Memory Jogger—A Pocket Guide of Tools for Continuous Improvement and Effective
Planning. 1994. 1st Edition. GOAL/QPC. Methuen, MA.
The Team Handbook—How To Use Teams To Improve Quality. 1992. 18th Printing. Joiner
Associates, Inc. Madison, WI.
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Course
Outline
FISH 221
Field Notebooks & Journals
Field Notebooks and Journals
INTRODUCTION
As natural resources, fisheries and/or GIS technicians, one of the most important tools you will
need to possess will be to observe and record what goes on around you in the natural world. Field
notes and journals are an important part of developing and improving your observation and datacollection skills over time, and you will be required to keep both a field notebook and journal this
entire quarter. Observations, insights, and key information you learn in this class will not only help
you in this class, but it will provide an excellent record for you to refer back to whenever needed.
You will develop record-keeping skills by keeping field notes and journals on a daily basis. Periodically, both your field notebooks and journals will be turned in to be reviewed by the instructor or
his/her assistant.
Use your text, “The Naturalist’s Field Journal,” as your guide for both format and content for your
field notebooks and journals (see course outline—text and references).
FIELD NOTEBOOKS
The field notebook is usually a Rite-In-The-Rain type (either the 3.5" x 5" size, or the spiral
bound). Both are designed to fit in your shirt or back pocket. Both types can be purchased for
about $3-4 at the college bookstore. You may fill up several notebooks during one quarter! Always
write in your notebooks using a #2 pencil or black waterproof ink. Most inks will “run-in-therain.” Remember too, that field notebooks always go with you into the field.
Use your “Rite-in-the-Rain” field notebooks to record your observations and measurements (when
field data forms are not provided). Also record questions and insights and make diagrams and
sketches whenever possible. At the end of the day, you should always transcribe your field notes
into your journal.
Items in your field notebooks should describe: who?, what?, when?, where?, why? and how?
1. Front Covers:
Some important information to include on the front cover of your field notebooks
includes:
• course number and course name
• a note that says, “If found, please return to: (your name, address and phone #)”
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Field
Notebooks
2. Inside the Front Cover:
Important information to write inside the front cover includes the following reminder
checklist:
•
•
•
•
•
•
•
•
•
•
•
•
•
date/time
weather observations
locations, landmarks
routes to and from your destinations
field notes from instructors or guest speakers (in the field)
mileage to and from destinations
plants, animals and habitats observed
questions and/or insights
qualitative observations based upon your senses of sight, smell, hearing, touch and taste
quantitative measurements (when field data sheets are not provided)
animal signs (tracks, scat, calls, etc.)
maps, drawings and sketches
who was with you on your data-collection trip?
NATURALIST’S JOURNAL
You will transcribe notes taken in the field into your journal. Unlike your field notebooks, your
journal does not go into the field.
Your naturalist’s journal consists of three major sections: The Journal, Species Account and Diary.
1. The Journal:
This first section is much like your field notebook. Much of the information you recorded
that same day in the field can be transcribed to this section of your journal. Things to
avoid in this section are personal feelings, emotions, and comments of a non-scientific
nature (save these for your Diary). Journal entries are a bit more formal than field
notebook entries (pay more attention to complete sentences, grammar and punctuation).
Journal entries can serve as legal evidence and are often bound and placed in library
collections.
Follow the examples in Herman’s text for both format and content, and you can’t go
wrong.
It goes into great detail as to the specific format and style of the field journal. You will be
required to learn and use this style. Consult Herman’s text whenever you are in doubt. His
book is based upon the Grinnell System of Journal Writing, which is a very disciplined type
of technical writing.
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Field
Notebooks
2.
Species Account:
The first time you see a particular species of a plant or animal, it should be noted in your
field notebook. Sketch it right on the spot! Describe the habitat you found it living in.
What other species of plant and animals were also living in this habitat? Describe what life
stage or season of the year you observed the plant or animal.
As you transcribe this new species to your Species Account Section of your journal,
describe in full detail what you observed the first time you noted it. This may take more
than one page. You may have to look up some scientific information or even key out the
species the first time you record it. As minimum, you should include:
•
•
•
•
•
•
•
•
•
•
•
Common name
Scientific name (genus and species)
Other taxonomic classifications (phylum, class, order, and family)
date/time of observation
location
habitat
season of the year
other plants and animals nearby
ecological relationships (interactions with other members of the observed
ecosystem)
physical, chemical, and biological characteristics (size, color, smell, sex, life stage,
texture, etc.)
sketch of what you observed
With the second and subsequent sightings, you only have to make general observations
such as date, time, location—place these entries directly below your previous entries for
this species. Draw additional sketches if the appearance is different than previous
observations (i.e., perhaps this eagle is an adult compared to the immature one you saw the
first time; or you now observe a salmonberry [Rubus spectabilis] bush in winter—without
leaves—as compared to the first observation in the spring of the year).
3. Diary:
The Diary section of your journal is where you can be the most creative. There are almost
no restrictions on what you can write about in this section. Many students like to express
their emotions and feelings about what they saw in the field that day, or how the class
itself is going for them (good or bad). It provides a chance to “vent some steam” if
necessary or “go nuts over something that really turned you on” that day. Remember that
your diary entries will be kept strictly confidential. No one but your instructor and
assistant will ever see what you write here. We often make comments in this section that
either compliment or disagree with your comments and/or thoughts. Please keep our
comments confidential too!
NOTE: Try to write equally in all three (3) sections every day you write in your journal.
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Field
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PROCEDURE
For this first exercise, you will have a chance to use all of your senses to observe a “bit of nature”
around you.
A. Field Scenario
You will choose a small portion (a few square meters) of the natural environment around you to
observe for this exercise. This involves sitting quietly in your chosen spot and recording in your field
notebooks all the required information (see the checklist you’ve put inside the front cover of your
notebook), plus everything else you see, hear, smell, touch, or taste within your “private space.” Pay
special attention to the living things you observe: Where are they?, what are they doing?, what
sounds do they make?, what do they look like? (provide a sketch), how big are they? (measure them),
what color are they? (NOTE: You should be equipped with a metric ruler and a hand lens). Also note the
physical and chemical environment: air temperature, weather, clouds, soil type, water conditions,
elevation, etc.
Be sure to observe, both close in and far away from your chosen location, for at least 90 minutes.
Record as much as you can during this time. (NOTE: This field notebook scenario was the “brainchild” of
Ms. Judy Moore, Biology Instructor, Yakima Valley Community College, Yakima, WA. It is included with her
permission).
B. Naturalist’s Field Journal Exercise
The same day (or night) after you make your initial field observations in your field notebooks, take
some time to transcribe your records equally in the three (3) sections of your journals. There should
be plenty of information to fill a number of pages in each section. Be sure to consult Steve
Herman’s text for format and content. You may also have to consult taxonomic keys for animals and
plants when you do your species accounts. (NOTE: Sketches should be your own, not a photocopy of a
picture from a book.)
Remember that in the Diary Section of your journal, it’s your chance to reflect on how this nature
observation exercise related to you as a person. How did you feel as you were sitting quietly observing and
writing? Does this bring up any memories of your past? How do you, or humans in general, fit into or relate to
nature today? This section of your journal will feel very much like you are writing an English paper—
you have “the freedom to explore and expound upon” your feelings, emotions, opinions and put
them into writing, very unlike most scientific or technical writing you will do. And it’s OK to take a
few notes in your field notebook to jog your memory later when you write them in more detail in
your diary.
LAB PRODUCTS
Turn in your field notebooks and journals.
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Field
Notebooks
FISH 221
Plankton Sampling & Analysis
Plankton Sampling and Analysis
INTRODUCTION
Plankton are defined as organisms suspended in a body of water which, because of their physical
characteristics or size, are incapable of sustained mobility in directions counter to water currents.
Most plankton are microscopic and of essentially neutral buoyancy. All of them drift with the
currents (APHA, 1985).
Plankton consist of both plants (phytoplankton) and animals (zooplankton), and complex
interrelationships exist among the various components of these groups. Chlorophyll-bearing plants
such as algae usually constitute the greatest portion of the biomass of the plankton. Phytoplankton use the energy of sunlight to metabolize inorganic nutrients and convert them to complex
organic materials (they are autotrophic). Zooplankton and other herbivores graze upon the
phytoplankton (they are heterotrophic), and, in turn, are preyed upon by other organisms (i.e.,
fish and shellfish, and amphibians), thus passing the stored energy along to larger and usually more
complex animals (among trophic levels). The heterotrophic animals in these food chains (or
more commonly, food webs) release organic materials to the aquatic ecosystem (through waste
products and when they die). These in turn are broken down into inorganic chemicals by bacteria
(also heterotrophs). These inorganic nutrients are released into the water and become useable
again by the primary producers (the phytoplankton). Thus we say, “energy flows and nutrients
cycle.”
Plankton may form the “base” of the food web in most aquatic ecosystems. Therefore data
concerning them may be particularly significant to the water quality technician or pollution
biologist. Phytoplankton “blooms” often cause extreme fluctuations of the dissolved oxygen in
lakes. Phytoplankton both respire (use oxygen) and release oxygen (through photosynthesis)
during any given 24-hour period. Large concentrations of plankton can cause offensive taste and
odor problems in water or in the fish that inhabit those waters. They can also be aesthetically
objectionable (causing the water to have a brownish, turbid appearance). The quantity of the
phytoplankton occurring at a particular sampling location depends upon many factors including
sampling depth, time of day, season of the year, nutrient content of the water, and presence of
toxic materials.
During the next two lab/field exercises, you will learn to collect and analyze both phytoplankton
and zooplankton from a lake. We will most likely be able to trace a plankton “bloom,” which
usually begins around early spring (April-May), and continues to increase until the nutrients in the
lake become limiting. Zooplankton “blooms” often lag behind the blooms of phytoplankton by
several days (or weeks). Since they are higher on the food pyramid than the phytoplankton, the
biomass (or energy content) will be roughly one-tenth (1/10) that of the phytoplankton. We will
sample and analyze both the phytoplankton and zooplankton on a weekly basis throughout this
quarter to see if these trophic relationships hold true.
15
Plankton
PROCEDURE
A. General Considerations
Before plankton samples are collected, a study design must be formulated. The objectives must be
clearly defined, and the scope of the study must remain within the limitations of available personnel,
time and money. Historical, biological, chemical and physical (especially hydrological) data should
be examined when planning a study. Of particular importance are data concerning volume of the
lake, flow, currents, prevailing wind direction, temperature, turbidity depth (light penetration), and a
complete chemical analysis of the inorganic nutrients at all depths in the lake. (NOTE: These
observations were made in a previous class). During the next two lab/field exercises, we will do some
preliminary sampling and analysis to ensure we have the methodology mastered. We will then
apply these techniques in later labs (e.g., lake studies).
We will locate our plankton sampling stations as near as possible to those selected for chemical,
physical and bacteriological (Biological Oxygen Demand and fecal coliform) sampling stations to
ensure maximum correlation of findings. We will establish a 100-meter surface transect by anchoring
buoys at the beginning and end of the transect. NOTE: Our sampling site was used by previous students,
assuring the availability of historical data that will lead to a better understanding of our results and provide
continuity to our current study.
B. Sampling
For sampling phytoplankton and zooplankton in the lake, we will use the protocol described in
American Public Health Association’s (APHA) Standard Methods for the Examination of Water and
Wastewater (1985), pp 1043-1083.
We will use either a fine mesh plankton net towed from our 12 ft. Alaskan boat or a water sampler
to collect our samples. Plankton nets are preferred to bottles and traps when plankton are few or
when only qualitative data are needed for analysis.
The maximum volume (V) of water is that which can be filtered through a net during an oblique
tow (the tow line is maintained at a pre-determined angle [usually 60o] during the entire tow).
V = p r2d where:
p = pi (3.14)
r = radius of the net orifice
d = distance (meters) the net is towed
After the sample has been collected, rinse the net with sufficient quantities of tap water (1-2
gallons) to ensure all plankton have been washed from the inside of the net into the attached glass
vial. Fill out a sample label and place it inside the vial. Add preservative (if required) and screw
on a lid. Run a replicate tow (if required). For today’s lab, live samples will be examined. Store
samples in an ice chest in the boat, and in a dark refrigerator in the lab. NOTE: Keep the sample
at ambient temperature and examine specimens as soon as possible after collection (if possible).
16
Plankton
C. Preservation
The most suitable phytoplankton preservative is Lugol’s Solution, which can be used for all forms
including flagellated phytoplankton.
D. Concentration or Dilution
The plankton contained in water samples often have to be concentrated (if too few are contained
in the sample), or conversely, diluted if necessary. You will determine if concentration or dilution
is necessary when you begin your analysis. Standard Methods gives exact instructions on pp 105455. It is important to record the amount of dilution or concentration used since these factors will
be used in our final calculations of plankton density. A Sedgewick-Rafter Counting Cell will be
discussed under Item F. below.
E. Microscopic Identification of Plankton
You will use both a compound and binocular microscope for identification of the phytoplankton
and zooplankton in your mixed sample. A depression slide works well if there are some macroscopic zooplankton in the sample. Use a drop of Protoslo to immobilize “fast track” zooplankton.
Use plankton keys supplied by the instructor to identify plankton. Try, at least, to key your specimens to common name, genus and species. Draw a sketch of each plankton on the sheets provided.
F. Plankton Counting
We will use the counting protocol specified in Standard Methods (pp 1057-1067). This method
includes calibrating and using a Whipple Micrometer Square (Figures 3a. and 3b.) in the ocular lens
system of our compound microscopes (Standard Methods, pp 1057-60), and a Sedgewick-Rafter
(S-R) counting cell (pp 1061-63).
When determining the number of plankton in 1.0ml (within the Sedgwick-Rafter slide) you can
practice two different techniques: 1) the “field” method and/or 2) the “strip” method.
G. Estimating Plankton Density In the Lake
Once we know how many phytoplankton are in 1.0 ml of our sample, we can multiply this times
the number of ml in our sample (assuming no dilution or concentration). We can then calculate the
volume of water sampled by our plankton net towed over the 100 meter transect (V [m3] = pr2d).
We know that the number of plankton in our sample came from the volume (V) sampled by our
plankton net. Since we know the estimated volume of the lake (from supplied data) we can divide
the total volume (V) of the lake by the volume of the plankton tow. We can then multiply our
sample number of plankton by the number of possible plankton tow volumes in the entire lake
and get a fair estimate of plankton number at this particular point in time. Congratulations—you
made it!
17
Plankton
LAB PRODUCTS
A. Be sure you wash plankton nets with fresh tap water thoroughly after each day’s use. Hang
nets in the specified drying area before stowing away. Wash and stow all other sampling
equipment and supplies away where they belong.
B. Make sure you have recorded your observations and measurements in your field notebook.
C. Complete your drawings and data sheets for both your qualitative observations (keying out
plankton) and your quantitative measurements (plankton counting, density calculations and
graphic plot of plankton).
D. Turn in your field notebooks, your drawings, data sheets and a 2-3 page, individually prepared
write-up of this lab/field exercise.
18
Plankton
Plankton Data Sheet
Names of Samplers:
Type of Sample:___________________
Location:
___________________
Subsurface tow: __________ft.
Length of Tow: _________m.
D ate /
Time
Tow #1
(Surfac e )
Tow #2
(Surfac e )
Tow #1
(Subsurfac e )
Tow #2
(Subsurfac e )
#Phtoplank ton
#Zooplank ton
Calculations: (Show work)
19
Plankton
Graphic Plot of Plankton Numbers
Lake ____________________ Months Sampled:______________________
Names of Samplers:
1020
1015
Logarithm
of
Plankton
Numbers
1010
105
101
#1
#2
#3
#4
#5
Start date
#7
#8
#9
End date
Sample Weeks
20
Plankton
#6
Specimens Collected
Name _______________________________ Section __________ Date ________
Spe c ime n N o.
D rawing
D ivision
Common N ame
Ge nus (if k nown)
Habitat
21
Plankton
NOTES FOR INSTRUCTORS
Be sure to read through this exercise and these notes fully before planning to implement this activity
with students, as some information needs to be researched and reviewed beforehand.
A. General Considerations
We typically use a sampling site used by previous students, assuring the availability of historical
data that will lead to a better understanding of our results and provide continuity to our current
study.
When you design the comprehensive lake study, you may want to use a grid network established by
Global Positioning Systems (GPS) and Geographic Information Systems (GIS). Also, you may
want to establish two parallel transects (replicates) if your statistical design calls for them.
B. Sampling
For sampling phytoplankton and zooplankton in the lake, use the protocol described in American
Public Health Association’s (APHA) Standard Methods for the Examination of Water and
Wastewater (1985), pp 1043-1083.
Once the sampling locations, depths and frequency have been determined, students should
prepare for sampling by labelling sample containers (glass vials) with sufficient information to
avoid confusion or error. Cut ¾" X 2" strips of Rite-In-The-Rain paper (20 strips per team) for
labels. On the label (in pencil) indicate date, sampling station, study area (lake name), type of
sample (plankton net), and depth (surface or subsurface). If subsurface sampling, describe how to
estimate the depth of the tow. If the samples are to be preserved, list the type of preservative and
the concentration on the label. Also; if replicate samples are taken, put the tow# on the label.
In their field notebook students should record the sample location, depth, type, date, time,
meteorological conditions, turbidity (secchi disk reading), water temperature, and other significant
observations. These field data may be invaluable when analytical results are interpreted, and often
help to explain unusual changes caused by the variable character of the aquatic environment
(APHA, 1985).
The maximum volume (V) of water is that which can be filtered through a net during an
oblique tow (the tow line is maintained at a pre-determined angle [usually 60o] during the entire
tow).
V = p r2d where:
p = pi (3.14)
r = radius of the net orifice
d = distance (meters) the net is towed
Since the cosine of 60o is 0.5, the depth of the tow can be estimated based upon the principle of
right triangles. For example, if the tow line length (the hypotenuse) is 50 ft., and a tow angle of
22
Plankton
•FOR INSTRUCTORS•
60o is maintained during the entire tow, then the depth is ½ of the length of the tow line (50 ft)
minus the distance of the top of the tow line to the surface of the water. Thus, if the distance of
the tow line above water is 4 ft., then the actual depth of the plankton net would be 21 feet.
C. Preservation
The most suitable phytoplankton preservative is Lugol’s Solution, which can be used for all forms
including flagellated phytoplankton. See Standard Methods (p. 1047) for preparing Lugol’s Solution.
Preserve zooplankton samples with 70% ethanol or 5% buffered formalin. Ethanol preservative is
preferred if the zooplankton are to be stained (pg. 1052, Standard Methods, 1985). Formalin may
be used for the first 48 hours of preservation with subsequent transfer to 70% ethanol.
D. Plankton Counting
Use the counting protocol specified in Standard Methods (pp 1057-1067). This method includes
calibrating and using a Whipple Micrometer Square (Figures 3a. and 3b.) in the ocular lens system
of a compound microscopes (Standard Methods, pp 1057-60), and a Sedgewick-Rafter (S-R)
counting cell (pp 1061-63). The S-R counting cell (Figure 2b.) is a device commonly used for
plankton counting because it is easily manipulated and produces reasonably reproducible results
when used with a Whipple disc. The S-R cell is 50mm long by 20 mm wide by 1mm deep. Thus,
the total area of the well is 1000 mm2 and the volume when filled w/sample is 1000 mm3 or 1.0
ml, or approximately 1.0 cm3. This will be very useful when we start to extrapolate the number of
plankton in 1.0 cc to the number in the column of water sampled by the net (# plankton/m3) and
finally, the number of plankton in the entire lake.
When determining the number of plankton in 1.0ml (within the Sedgwick-Rafter slide) you can
practice two different techniques: 1) the “field” method and/or 2) the “strip” method. In the first,
10 randomly chosen fields of view (within the calibrated Whipple disc) are counted for phytoplankton and zooplankton. An average #/field is then calculated. Using the “strip” method, 34 full-length strips across the length of the S-R slide are counted. These are then averaged to get
#/strip. You can then calculate the number of “field,” or “strips,” per S-R slide (based upon our
original micrometer calculation for the Whipple disc). Then multiply or divide number of cells per
milliliter by a correction factor to adjust for sample dilution or concentration. Standard Methods
outlines the calculations to find # of plankton per ml. of sample for fields and strips on pp 106263.
•FOR INSTRUCTORS•
23
Plankton
REFERENCES
American Public Health Association. 1985. Standard Methods for the Examination of Water and
Wastewater. 16th edition. WA, D.C., 1268 pp.
Cupp, E.E. 1943. Marine plankton diatoms of the west coast of North America. Bull. Script
Inst. Oceanogr. 5:1
Davis, C.C. 1955. The marine and fresh water plankton. Michigan State University Press, East
Lansing, MI.
Hof, F.H. and T.W. Snell. 1997. Plankton culture manual. 4th edition. Florida Aqua Farms, Inc.
Dade City, Florida.
Needham, J.G. and P.R. Needham. 1962. A guide to the study of fresh-water biology. 5th edition.
Holden-Day, Inc. San Francisco, CA.
Prescott, G.W. 1978. How to know the fresh water algae. 3rd Ed. Wm C. Brown Co., Dubuque,
Iowa.
Reid, G.K. 1967. Pond life—a guide to common plants and animals of North American ponds
and lakes. Golden Press, New York, New York.
Whipple, G.C., G.M. Fair and M.C. Whipple 1927. The microscopy of drinking water. John
Wiley and Sons, New York, N.Y.
24
Plankton
•FOR INSTRUCTORS•
FISH 221
Biometrics — Terminology & Problems
Biometrics—Terminology and Problems
Biometrics is an important tool used to study natural resource-related entities—whether they be
trees, wildlife, invertebrates, or fish. The following provides an overview of concepts and related
sample problems.
TERMINOLOGY
Statistics
For our purposes, this may be defined as a collection of methods which have been
developed for handling numerical data.
Biometrics
The application of statistical methods to the solution of biological problems.
Experiment
Any scientific endeavor where observations or measurements are made in order to
draw inferences about the real world. Some experiments are rigidly controlled,
whereas others can take the form of a study, survey or field collection.
Observation A record representing some property or characteristic of a real world object.
Measurement is used in place of observation and implies a quantified observation.
Character of That observation or measurement related to the purpose of the study. The
Interest
character of interest is the characteristic being observed or measured; the
measurement is recorded, analyzed and interpreted to draw an inference about
the real world.
Experimental The experimental unit is the object upon which the observation is made—it could
Unit and
be a single fish, an aquarium full of fish, or a lake full of fish. It must be clearly
Universe
defined so as to restrict measurements to only those units of interest to the study.
The universe is the set of all experimental units of interest in the study.
Population
and Sample
Population refers to the set of values for the characteristic of interest for the
entire group of experimental units about which the inferences are to be made (the
universe). Observations are not taken for all possible experimental units—only a
sample is taken. A sample is a set of observations, usually only a small fraction
of the total number that will be taken. Thus, samples represent a portion of the
real world that has been selected for measurement.
Statistic and
Parameter
The sample statistic is an estimator of the population parameter, and there is a
certain degree of variation among samples. When trying to characterize a
population, we cannot obtain a perfect representation of that population by only
25
Biometrics
looking at a sample. For example, if we want to know about the population
mean, we take an adequate sized sample and determine the sample mean. The
population mean is a parameter. The sample mean is a statistic. The statistic is
related to the parameter in the same way the sample is related to the population.
Hence, we speak of population parameters and sample statistics.
Frequency
Indicates how many times a particular score occurs in a collection of data.
Frequency
A tabular arrangement of data, ranked in ascending or descending order of
Table
magnitude, together with the corresponding frequencies.
Frequency
Histogram
A set of rectangles having bases on a horizontal axis with centers at the given
scores and heights equal to the corresponding frequencies (see figure 1 below).
Frequency
Polygon
A line graph of frequencies plotted against scores (can be obtained by connecting
midpoints of tops of rectangles in the frequency histogram).
To illustrate the definitions above, consider a study in which chloride levels in a lake are to be
determined. Twelve water samples are taken and chloride levels measured in µg/L. These
measurements are given below:
100
100
100
101
100
98
99
99
101
101
102
102
These measurements may be arranged in a frequency table that represents how often each
measurement occurred:
F r e q u e n c y Ta b l e
26
Biometrics
Chl ori d e ( µg/ L)
F re q u e nc y
98
1
99
2
100
4
101
3
102
2
This table can then be used to generate a frequency histogram and frequency polygon as
follows:
Frequency
4
3
2
1
98
99
100
101
102
Chloride [g/L (ppb)]
Figure 1. Frequency histogram for chloride concentrations (µg/L or parts per billion).
A. Measures of Central Tendency
Central Tendency
The tendency of values to cluster about a particular value in a distribution
of data.
Mode
The value which occurs most frequently. In the example above,Mode = 100.…g / L
Median
Midpoint of a distribution of scores. It is most commonly used when the
distribution is skewed or asymmetric.
If there are an odd number of observations, n, the median is x (n+1)/2 where x (n+1)/2
represents the ((n+1)/2) th value in the frequency distribution.
If there are an even numbeer of observations, the median is (x n/2 +x (n/2+1) )/2
Median=
Mean
x n/2 +x (n/2+1)
2
=
x 6 +x 7 100+100
=
=100…g/L
2
2
Arithmetic average for all the values in the sample distribution, denoted byx.
The formula for calculating the sample mean is:
x=
x1+x2+x3+...+xn
n
n
x=
x=
∑x
i
i=1
n
∑x
n
where there are n observations in the sample
Mean=x=
∑ x = 98+2(99)+4(100)+3(101)+2(102) =100.25…g/L
n
12
27
Biometrics
B. Measure of Dispersion
Dispersion
Spread or variability of observations in a distribution.
Range
The difference between the higher value and the lowest value.
Range = 102 - 98 µ g / L = 4 µ g / L
Average
Deviation
The sum of the deviation of the values from their mean, without regard to the
sign, divided by the total number of values. The formula for calculating the
average deviation (AD) is:
AD =
∑ x−x
n
nx
x
x-x
nx x - x
1
2
4
3
2
98
99
100
101
102
2.25
1.25
.25
.75
1.75
2.25
2.50
1.00
2.25
3.50
11.50
Variance
AD =
11.5
µg / L
12
2
∑ (x − x )
=
n −1
2
where there are n observations
nx
x
x−x
(x − x )
nx ( x - x )
1
2
4
3
2
98
99
100
101
102
-2.25
-1.25
-.25
+.75
+1.75
5.06
1.56
.06
.56
3.06
5.06
3.12
.24
1.68
6.12
16.22
s2 =
Note: Using the “absolute value”
of the difference between each value
of x and the mean xwhen solving for
average deviation, is one way of
disregarding the “+” or “-” signs.
The sum of the squares of the deviation of the values from their mean
divided by the total number of observations minus 1.
s
28
Biometrics
where there are n observations.
2
2
Note: By squaring the difference
between each value of x and the
mean x in determining the variance,
we can disregard the “+” or “-” signs.
NOTE: Using the divisor (n-1) instead of n in computing s2 seems puzzling at first. It can be
shown that dividing by n-1 provides an unbiased estimate of the population variance,
whereas dividing by n produces estimates that are a bit too small (dividing by n tends to
underestimate the population variance).
Standard Deviation
The square root of the variance s=
( )
Σ x-x
2
n-1
where there are n observations
The formula adapted for the hand calculator is:
s2
(∑ x)
∑ x2 n where there are n observations
s=
n-1
2
_x
_ _x
nx x
_,
nx
x
1
98
98
9604
9604
2
99
198
9801
19602
4
100
400
10 0 0 0
40000
3
101
303
10 2 0 1
30603
2
102
202
10404
20808
Totals
1203
x
2
_jjjj2j
nxx
120617
12032
12
11
120617s=
s=
120617-120601
11
s=
16
=1.21µ g/L
11
29
Biometrics
When using a hand-held calculator, the following calculation method may be helpful:
Subtracting a constant from each score in the distribution does not affect the variance or the
standard deviation of the distribution. We can simplify the computations by first subtracting 100
from each score in the distribution, thus simplifying our calculations considerably.
_,
nx
_ _x
_x
nx x
x
_ j j j j2j
n xx
1
-2
-2
4
4
2
-1
-2
1
2
4
0
0
0
0
3
1
3
1
3
2
2
4
4
8
Tota ls
s =
3
Σx
2
s2 =
s=
( Σx )
-
17
2
n
n -1
2
Relative Standard
Deviation
x
2
( 3)
17 -
2
12 = 16.25 = 1.48 µ g / L
11
11
Σxi
2
( Σxi )
n
n -1
2
= 1.48 = 1.22 µ g / L
The standard deviation expressed as a fraction of the mean.
The relative standard deviation is often expressed as a percent
and referred to as the coefficient of variation:
c=
(s)
(100)
x
The relative standard deviation is particularly helpful when comparing the precision of a number of determinations on a given substance at different levels of concentrations.
30
Biometrics
Statisticians often represent data with theoretical curves which are
smoother than frequency polygons, obtained from experiments in
real life. However, frequency distribution or frequency polygons of
experimental data often approximate a mathematical function called
the “normal” distribution curve.
Frequency
Normal Distribution
Curve
Quantity Measured
Figure 2. Normal Distribution Curve
In figure 3 the frequency polygon for chloride in water is a fairly good approximation of the
“normal” curve.
Frequency
4
3
2
1
97
98
99
100
101
102
Chloride m g/L
103
Figure 3. Normal Distribution Frequency Polygon
31
Biometrics
If a frequency distribution is a good approximation of the normal curve, we can use some facts
about the normal curve to give us information about the frequency distribution. Figure 4 shows us
the normal distribution in terms of the mean (µ), and the standard deviation (σ), and gives the
percent of area under the curve between certain points.
68%
95%
99%
-3
-2
-1
+1
+2
+3
Figure 4. Normal Distribution Curve.
Figure 5 shows the frequency polygon obtained from the chloride determinations in terms of the
mean (x), and the standard deviation (s), and gives the percent of area under the curve between
certain points.
Frequency
4
3
2
75%
1
100%
-2s
-1s
98
99
_
x
100.25
+1s
101
102
+2s
Figure 5. Comparison of Normal Curve and Frequency Polygon
Comparison of Figures 4 and 5 shows that we are justified in saying that the frequency distribution
approximates the normal distribution. Specifically then, assuming a normal distribution we would
expect 68% of the observations to lie within ± 1σ from the population mean when in fact, 75% of
the observations were within ± 1s from the sample mean. Likewise assuming a normal distribution
we would expect 95% of the observations to lie within ± 2δ from the population mean when in
fact 100% of the observations were within ± 2s from the sample mean. In both cases the
observed percentages are reasonably close to the expected percentages. Tests exist for determining
whether or not a frequency distribution might reasonably be assumed to approximate the normal
distribution.
32
Biometrics
Biometrics Problem
Codfish Length—Frequency Distribution
Given the following data collected on lengths of codfish, answer the questions.
Le ngt h
F re ( c m) q u e nc y
( n)
(l)
l
n
l
n
l
n
l
n
l
n
25
2
39
18
53
15
67
--
81
--
95
--
26
7
40
15
54
8
68
--
82
2
96
--
27
8
41
13
55
6
69
--
83
1
97
--
28
9
42
13
56
11
70
1
84
--
98
--
29
13
43
19
57
7
71
1
85
--
99
--
30
12
44
19
58
4
72
--
86
1
100
--
31
9
45
21
59
5
73
1
87
1
101
--
32
15
46
13
60
1
74
--
88
--
102
1
33
7
47
19
61
2
75
--
89
--
103
--
34
7
48
21
62
1
76
--
90
--
104
--
35
5
49
8
63
2
77
1
91
--
105
--
36
12
50
22
64
--
78
1
92
1
37
13
51
18
65
--
79
--
93
--
38
16
52
18
66
2
80
1
94
-Tota l 4 4 9 =n
1. Plot the following data for length-frequency distribution of North Pacific black cod as
frequency polygons and as histograms, first at 1 cm intervals, then at coarser groupings of 5
cm, 10 cm and 20 cm intervals.
a. Compare the effect of using different base points for grouping (e.g., for the 5 cm group,
use 25-29, 30-34, etc., or 27-31, 32-36, etc.). Use a 5 cm grouping only on this.
b. The plots should be made on the same effective scale—(e.g., when using 5 cm intervals,
the frequency in each interval will be about 5 times that when using 1 cm intervals so
that the frequency should be reduced accordingly (i.e., for 1 cm intervals, 1 fish equals
33
Biometrics
1 unit on the graph paper; for 5 cm intervals, 1 fish equals 1/5 unit on graph paper;
for 10 cm intervals, 1 fish equals1/10 unit on the graph paper, and 1/20 unit for 20 cm
intervals).
c. The length (along the x-axis of your graph) should not be altered. Plotted this way, the
various polygons and histograms should be nearly identical in appearance.
This example will illustrate the problem of the correct choice of class interval.
What appears to be the best grouping to use? Why?
2. You will find that many frequency distributions will, when plotted, appear as single peaks with
more or less extensive tails above and below the peak.
a. Where does the peak (or peaks) appear in your distribution?
b. One or more quantities may be used to define the position of the peak of the distribution. The most commonly used is the arithmetic (or sample) mean, or more simply,
just the mean. Calculate the mean by the longhand method described above (hand
calculators are permitted). Show your work.
Formula for calculating sample mean:
n
x=
∑x
i
i=1
n
(where there are “n” observations in the sample)
NOTE: The numbers above and below the Σ (summation) symbol tell you what
quantities to add up: Sum of xi when i = 1, 2, 3… nth (or last observation, in
this case, the 449th observation).
34
Biometrics
c. Calculate the mode (the value at which the actual peak, or peaks, occur) and the median,
or 50% point, which is the value such that half the individuals in the population have
values less than the median, and half greater (use formulas given above). In most
distributions, the median will lie between the mean and the mode, being rather close to the
mean.
d. Where does your median lie?
3. a. Calculate the variance for these data.
b. The sample standard deviation is an average value of the deviation from the mean.
It is calculated by taking the square root of the sample variance ( s 2 ). Calculate
the sample standard deviation for this data.
4. Are your data normally distributed? How do you know?
5. Construct a distribution curve that shows the distribution of your sample data.
[This outline was prepared by L.A. Lederer, Statistician, Analytical Reference Service, Training
Program, NCUIH, SEC. The original document and date where published is unknown.]
35
Biometrics
FISH 221
Stream Analysis—Electroshocking
Stream Analysis—Electroshocking
INTRODUCTION
A.
General Information
Electrofishing or electroshocking is the use of electricity to capture fish. Electroshockers are
particularly useful in areas where uneven bottoms, fast flowing water, or obstructions (i.e., large
amounts of wood debris) are present and make conventional collecting techniques difficult or
impossible (APHA, Standard Methods for the Examination of Water and Wastewater, 1985).
Electroshockers are non-selective in terms of size and type of fish caught; however, in the streams
or ponds we will sample, we will be interested in all fish species living in these waters.
PROCEDURE
A. Operating Instructions
We will follow the operating instructions specified for the backpack electroshocker built by SmithRoot, Inc. (Vancouver, WA) commonly used for electrofishing in streams. This battery-powered
unit delivers 150 minutes of continuous power before recharging is required. This unit delivers both
D.C. and pulsed D.C. at 15-120 HZ and weighs 66 kg. The operating instructions are found both in
the manual and on the instrument itself.
B. Safety Aspects
Electrofishing is hazardous work! The batteries used in electrofishing provide enough energy to electrocute a person. The current that passes through the body and vital organs is particularly dangerous
(Reynolds, 1996). We will read safety precautions over as a class before we attempt any
electroshocking.
Before any electroshocking begins, both while learning these procedures and once we use the
technique for our field study, be aware of and practice the following safety procedures (Reynolds,
1996):
36
Electroshocking
1.
Never electroshock alone!
2.
Avoid more than 2-3 samplers per team!
3.
Have at least one experienced biologist or technician per team. That person must actively
supervise.
4.
Avoid operating near bystanders, pets or livestock in the creek or near the shore.
5.
Electroshocking should proceed slowly and deliberately—avoid chasing fish.
6.
Resist the urge to hand capture a stunned fish—the net person will use a net. A missed
fish is better than a “shocking experience.”
7.
Always shut down the power source when equipment changes, repairs or any other
non-routine situations arise.
8.
Rest often enough to avoid fatigue.
9.
Reread the procedures of this lab/field exercise before each and every sampling trip.
(We have enclosed these safety procedures in plastic laminate and have attached it to
the aluminum carrying case.)
NOTE: Team supervisors must ensure all team members have read and understand these procedures.
C. Field Sampling Scenario
Prior to beginning work, all students will be required to read and sign the attached Acknowledgment of Electrofishing Orientation.
1.
Teams of 2-3 students will sample a 50-100 meter stretch of _____ Creek. Proceed as
follows:
a.
Perform a conductivity measurement, and take a stream temperature. Record data
in your field notebooks.
b.
The person with the backpack shocker and the net person will be dressed in hip
boots or waders, elbow-length rubber gloves and raingear (if raining). Polarized
sunglasses are extremely helpful. The netter should carry a fiberglass-handled
shallow bag net. The person on shore should carry a partially-full, 5-gallon bucket
of stream water (or water with MS-222, if fish are to be narcotized before examination). A second partially-full, 5-gallon bucket can be used as a recovery tank.
NOTE: Fish should be fully recovered before release back to the stream.
The stream bank person will key out each fish to common name, genus and species;
determine life stage, length, and sex; and its general health and condition.
c.
Record all data on the field data sheet. This data sheet has been designed to allow a
tally of each species to be kept.
d.
The next sampling team will continue its 50-100 meter survey upstream of the first
beginning where the first team ended, and so on, with the third and fourth teams. 37
Electroshocking
LAB PRODUCTS
A.
Following this field exercise, all equipment, materials and supplies will be packed away and
stored where directed. Be sure to put all batteries on the battery charger.
B.
Ensure that all routine data (weather observations, etc.) and fish data have been recorded in
your field notebooks and field data sheets.
C.
Submit a 3-5 page, team-prepared, word processed or typewritten summary of this lab/field
exercise. Your summary should have a title page, table of contents, 2-3 pages of text
(including methods and materials, results—including tables and/or graphs), a discussion, and
references cited. An appendix can be included if you choose to include your raw data,
operating instructions, safety precautions, etc.
38
Electroshocking
ACKNOWLEDGMENT OF ELECTROFISHING ORIENTATION
I have received instruction and orientation about electrofishing from my teacher. As a result, I
understand and accept the following conditions:
1.
Electrofishing (EF) is an inherently hazardous activity in which safety is the primary concern.
The electrical energy used in EF is sufficient to cause death by electrocution.
2.
During operation, it is critical to avoid contact with the electrodes and surrounding water. The
EF field is most intense near the electrodes and can extend 5-10 m outward.
3.
The electrodes are energized by the power source, a generator or battery, and controlled by
safety switches; these switches must remain off until the signal is given to begin EF.
4.
The power source has a main switch that must be turned off immediately if an emergency
occurs.
5.
The electrodes are usually metal probes suspended in the water. If direct current is used from a
boat, the anodes (+) are in front of the boat to catch fish and the cathodes (–) may be
suspended from the sides; both can produce electroshock. When a metal boat is the cathode,
the boat is safe—as long as all metal surfaces inside it are connected to the hull.
6.
Moveable anodes on a boat are dangerous, especially on metal boats. All electrodes on a
conventional EF boat should be in the fixed position during operation.
7.
Dry skin and clothing are good protection against electroshock. The body should be fully
clothed during EF. Rubber knee boots are minimal foot protection, as are rubber gloves for the
hands. A personal flotation device must be worn when the water is considered swift, cold, or
deep. Ear protection is necessary for those working near the generator.
8.
At least two members of the EF crew must have knowledge of CPR and first aid. A first aid
kit and, in an EF boat, a fire extinguisher, must be within immediate reach during an operation.
Electroshock can cause heart fibrillation or respiratory arrest; CPR can cure only the latter.
The EF crew must know the location of the nearest defibrillation unit.
9.
A communication system, particularly hand signals, must be available to all members of an EF
crew. When multiple anodes are used in a portable EF operation, the buddy system must be
used. Above all, NEVER OPERATE ALONE.
10. Stunned fish should be removed from the EF field as soon as possible and not subjected to
continuous electroshock by being held in the dip net. Using the anode as a dip net is unhealthy
for fish and people—and should be avoided.
11. An EF operation should proceed slowly and carefully; avoid chasing fish and other sudden
maneuvers. Night activities require bright, bow-mounted headlights. Operations should cease
during lightning or thunderstorms; use discretion during rain. Avoid EF too close to bystanders, pets or livestock.
39
Electroshocking
12. All EF crew members must know who their leader is and recognize his or her authority as final
in operational decisions. However, every crew member has the right to ask questions and to
express concern about any safety aspect of an EF operation. A crew member has the right to
decline participation in an EF operation, without fear of employer recrimination, if he or she
feels unsafe in such participation.
_____________________________________
Student Signature
_______________
Date
I have discussed the above-named conditions with the employee and am satisfied that
he or she understands them.
_____________________________________
Teacher Signature
________________
Date
Adapted from a form used for employers and employees; Reynolds (1995), with permission. In
Murphy B.R., and D.W. Willis. 1996. Fisheries Techniques. American Fisheries Society.
40
Electroshocking
Electrofishing Data Sheet
Names of Samplers: ______________________
______________________
______________________
______________________
Stream Location: ___________________
D ate /
Tim e
Com m on
N am e
G e nus a nd
Spe c ie s
Stream Reach #_________
Se x
( M /F)
For k
Le ngth
#Spe c im e ns
(tally )
General Observations:
41
Electroshocking
NOTES FOR INSTRUCTORS
Read the lab carefully, follow all safety protocols, and explain the following to students prior to
doing this lab.
See APHA, Standard Methods for the Examination of Water and Wastewater (1985); and Fisheries
Techniques 2nd Edition (1996) [devotes an entire chapter (pp 221-253)] for more about the art and
science of electroshocking. Anyone interested in achieving mastery of this technique should
spend considerable time reading and studying these references.
Electroshocking—Theory of Operation
An electric field in the water is produced by passing a current between two submersed electrodes or
between an electrode and the ground. We use a newer style electroshocker that has only one
electrode and a trailing cable that goes to ground (the water).
Depending upon their design, these electroshockers will produce either an alternating current
(AC), direct current (DC), or in some cases both (a current inverter is required to produce both).
AC stuns fish in its field, allowing them to be dipped from the water, whereas DC induces galvanotaxis—so that fish move toward one of the poles (the wand that contains the positive electrode).
Once they swim into the electric field created by the anode (wand) and cathode (ground wire), they
are stunned and can be netted. Newer DC electrofishers are usually battery powered. They are
particularly effective in turbid water or in waters with numerous obstructions or heavy vegetation
(since they tend to draw fish toward the electrode).
NOTE: AC devices aren’t used much anymore since they are more likely to kill or injure fish. Environmental
factors such as water hardness and available electrolytes can impact the effectiveness of electroshocking, and
this is why we always measure the conductivity of the waters we’re sampling.
Field Theory
As mentioned previously, the electrodes deliver voltage and current in the water to form a threedimensional electrical field. The current flows between electrodes of opposite polarity (i.e., via flux
lines) and voltage surrounds each electrode at right angles to the flux lines (i.e., equipotential lines).
The field is non-homogeneous and weakens with distance from the electrode as energy dissipates in
the water (Reynolds, 1996). The three parameters that apply to the electroshocker’s circuitvoltage, current and resistance-differ at various points in the water. Current density is the
current that flows through a 1-cm2 plane (of water). Voltage gradient is the change in voltage
over a 1-cm distance. Resistivity is a measure of the water’s resistance (to carrying a current in
the water). Water with an extremely high resistivity (i.e., deionized water) will be extremely poor
at conducting a current. The inverse of resistivity is conductivity and is measured in µmhos/cm.
Therefore, it is important to have some trace minerals in sampling waters in order for the
electroshocker to be effective (the water must be able to conduct a current with a high enough
42
Electroshocking
•FOR INSTRUCTORS•
voltage to attract or stun the fish). Brackish water may contain too many minerals (electrolytes).
Distilled water has very low conductivity (0.5 - 3.0 µmhos/cm). The conductivity of most freshwater bodies is between 50-1,500 µmhos/cm. On average, seawater is 500 times more conductive
than fresh water (Reynolds, 1996).
Electroshocking Effects
Fish behavior in an electric field depends on the nature and intensity of the wave form applied. In
any electrical field, a threshold of intensity must be reached to achieve a given behavioral response
(Reynolds, 1996). Lamarque (1990) offers this review of effects on fish of an electric field:
“In responding to AC, a fish tends to assume a position perpendicular to the electric current (flux
lines) thereby minimizing voltage gradient in its body. It may undulate in attempting rhythm with
the AC cycle, exhibiting oscillotaxis (forced movement without orientation, a “thrashing” motion).
At higher field intensity, tetany, or muscle contraction, occurs and the fish is immobilized. Fish
responses are less predictable with AC than DC. Too much intensity can and will kill the fish.”
In a DC field, a fish typically turns toward the anode and exhibits electrotaxis (forces swimming
towards an electrode). As the fish nears the anode, a new threshold causes narcosis (muscle realization) and loss of equilibrium. While under narcosis, the fish may continue to swim, upside down,
toward the anode. Tetany is achieved near the anode. Continuous DC elicits taxis and narcosis in
fish if the appropriate threshold is reached. The effects of pulsed DC are less predictable. Depending upon fish size and species, taxis and narcosis may not occur. Continuous DC requires more
power than does pulsed DC.
The importance of electrofishing-induced stress is often unrecognized by the technician or biologist
because captured fish may appear normal. However, the stress syndrome results in an abnormal
physiological state including a reduced respiratory level and acidosis. This stress may take hours
and/or days for complete recovery. Recovering fish are also more susceptible to predation, less
competitive and unable to feed. Wild fish require a longer period to recover than do hatchery fish
(Mesa and Schreck, 1989).
Biological Factors Related to Electroshocking
Electrofishing tends to be more effective for some fish than others. Fish with fine scales (i.e.,
salmonids) tend to be more vulnerable than those with thick scales (i.e., ciprinids). Fish that inhabit
shallower shoreline waters are also more susceptible to electroshocking. Electrofishing is selective
for larger fish of a species. For example, adult salmon require a much lower voltage setting (about
one tenth) than juvenile salmon. This may also be influenced by the territorial behavior of the adult
salmon.
•FOR INSTRUCTORS•
43
Electroshocking
REFERENCES
American Public Health Association. 1985. Standard methods for the examination of water and wastewater.
16th Edition. Washington, D.C. 1268 pp.
Hart, J.L. 1973. Pacific fishes of Canada. Bulletin 180. Canadian Government Publishing Centre,
Ottawa, Canada. ISBN 0-660-10459-8.
Lamarque, P. 1990. Electrofishing of fish in electric fields. Pages 4-33 in Cowx and Lamargue (1990).
Lee, D.S., C.R. Gilbert, C.H. Hocutt, R.E. Jenkins, D/E. McCallister, and J.K. Stauffer, Fr. 1980.
Atlas of North American freshwater fishes. Publ. No. 1980-12, North Carolina Biological Survey.
McClane, A.J. 1974. Freshwater fishes of North America. Holt, Reibart and Winston. New York, New
York.
Mesa, M.G. and C.B. Schreck. 1989. Electrofishing mark-capture and depletion mythologies evoke behavioral
and physiological changes on cutthroat trout. Transactions of American Fisheries Society. 118:644-658.
Marrow, J.E. 1980. The freshwater fishes of Alaska. Alaska Northwest Publishing Co. Anchorage, AK.
Murphy, B.R. and D.W. Willis, editors. 1996. Fisheries techniques, 2nd edition. American Fisheries
Society. Bethesda, Maryland.
Reynolds, J.B. 1996. Electrofishing. (pp 21-2253) In B.R. Murphy and D.W. Willis, editors. Fisheries
Techniques, 2nd edition. American Fisheries Society. Bethesda, Maryland.
Scott, W.B. and E.J. Crossman. 1973. Bulletin 184. Freshwater fishes of Canada. Canadian Government Publishing Centre. Ottawa, Canada. ISBN 0-660-10239-0.
44
Electroshocking
FISH 221
Stream Analysis Discharge
Stream Analysis—Discharge
INTRODUCTION
Measuring discharge, or water velocity passing a cross section per unit of time, is a task performed by natural resource technicians as part of establishing permanent reference sites for
gathering data about streams and rivers. The ability to accurately make and replicate stream
channel measurements over a period of years, and through changes in personnel, is vital. We will
use the procedure described on pages 44-48 in the U.S. Forest Service’s Stream Channel Reference
Sites: An Illustrated Guide to Field Techniques (April 1994) as our guide to performing this field
test.
Stream discharge (Q) is the volume of water passing a cross-section of a stream per unit of
time and is expressed as cubic feet per second (cfs). Discharge is simply velocity times cross
sectional area (Q = VA). Cross-sectional area (A) is determined by stretching a tape across the
channel to measure the distance (stream width) at the cross-section location. Depth is measured
with a calibrated rod. Area = depth X width in small increments across the channel.
We will use a Marsh-McBirney current meter for our velocity measurements. We will also learn the
float method for making velocity measurements when a flow meter is not available or when time is
limited.
EQUIPMENT AND SUPPLIES
(Flow Meter and Float Method)
•
•
•
•
•
•
•
•
•
•
•
•
Marsh-McBirney flow meter
spare battery for flow meter
field notebook and/or field data sheets
clipboard
pencils
fiberglass cloth measuring tape (min. length 25 ft.)
hand calculator
chest waders or hip boots
raingear
polarized sunglasses (optional)
stopwatch or wrist watch with second hand
5-10 floats (orange peels or water-soaked blocks of wood)
45
Discharge
PROCEDURE
Current Velocity Measurements (Marsh-McBirney Flow Meter)
Procedures for this exercise are found in the Meter manual. Figure 58 shows the exact types of
data you should enter in your field notebooks (or on your field data sheets).
Computing Discharge
We will calculate sample measurements and calculations in the laboratory before we go to the
stream. Record your field data on the data sheets provided.
Float Method for Current Velocity
The float method is a simpler way to estimate discharge provided velocity has been previously
measured and cross-sectional area calculated. Equipment for the float method measurement is
simple: a measuring tape, timer (a stop watch or digital watch) and 5-10 floats. For floats, use an
orange peel, a water-soaked block of wood, or other natural material that sinks at least halfway
into the water, is visible from shore, won’t be moved by wind and is expendable and non-polluting.
NOTES: Previously measured cross sectional area (from midsection method used for flow meter
calculations) can be used to find discharge (Q = VA), or if previous area data do not exist, an
average cross sectional area can be calculated by averaging 3-4 width and depth measurements at
the beginning, end, and along the channel segment being used.
LAB PRODUCTS
Students should switch roles as note-taker and meter reader after the first measurements have been
made (move to a new station 10 feet or so downstream for the second sampling). Each team of
two students will turn in a one-page report that describes these two methods for measuring discharge. Include your two field data sheets as enclosures to your report. Be sure to discuss your
results in your report. Turn in a one-page team report at the beginning of the next lab/field period.
Be sure that the names of both the note taker and meter reader appear on these forms.
REFERENCE
U.S. Forest Service, 1994. Stream channel reference sites: An illustrated guide to field techniques.
pp 44-48.
46
Discharge
Date & Time:
Location:
Note Taker:
Cros s S e c t i on
Me a s u re me nt
Discharge Field Data Sheet
Weather Observations:
Meter/Tape Reader:
Discharge Measurements (Marsh-McBirney Flow Meter)
Ta p e
Di s t a nc e ( ft )
Wi dt h
( ft )
De pt h
( ft )
Ve l o c i t y
( ft /s e c )
A re a
2
( ft )
Q= Di s c ha rge
( c fs )
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
(add al l Q's from s ub-s e ctions to ge t total dis charge )
Total Dis charge
=cfs
47
Discharge
Discharge Measurement Calculations
(Float Method)
Average Depth (Average of 3-4 measurements) =
_________________ ft.
Average Channel Width (Average of 3-4 measurements) =
_________________ ft.
(A) Area = Average depth X Average width =
__________________ ft.2
(V) Velocity = Distance (ft.)
Avg. Time (sec.)
Distance = _________________ft.
Time =_________________sec.
_________________sec.
_________________sec.
_________________sec.
_________________sec.
_________________sec.
Average Time = _________________sec.
V = Distance (ft) = Distance (ft.)
Avg. Time (sec.)
Adjusted V = V X *Coefficient for stream bottom resistance = __________ft/sec.
Q = AV (adjusted) = ________________cfs
*1. Coefficients for stream bottom resistance:
a. loose gravel, rocks = 0.8
b. smooth mud, sand, hardpan = 0.9
c. culvert = 1.0
2. Rate of flow (Q) X Correction factor for small streams (1.3) = __________cfs
Show your calculations below:
48
Discharge
49
Discharge
Tape
D istanc e (ft.)
Width
(ft.)
D e pth
(ft.)
Ve loc ity
(ft./se c .)
Total c fs:
A r e a (ft.2)
Q D isc har ge
(c fs)
Q=AV=_____ft2 x
_____ ft./se c . =_____c fs
A vg. Tim e (se c .)
V=D istanc e (ft.) X CF (dr ag) = ft./se c .
Tim e :
se c .__________
se c .__________
se c .__________
se c .__________
se c .__________
A vg. Tim e : se c .__________
D istanc e :__________
(V) Ve loc ity :
(ty pe of float use d:_______________________________)
(A ) A r e a:__________
Channe l Width:__________
A ve r age D e pth (Estim ate d):__________
D isc har ge Using Float Me thod
Meter Used: Marsh McBirney
Meter Reader:
Location:
WATER DISCHARGE (VELOCITY)
Comments:
Date/Time:
Method: Flow meter Measurements @ X-Sections
Note Taker:
FISH 221
Macroinvertebrate Sampling
Macroinvertebrate Sampling
The Surber Sampler, Ponar Dredge/
Grab, and Multiple-Plate Sampler
NOTE: This exercise will take 2 lab sessions to complete.
INTRODUCTION
A. Surber Sampler
Quantitative samples of invertebrates from shallow streams have traditionally been taken by using a
Surber sampler. A Surber sampler is a fine mesh net (with sides and a bag) attached to a hinged
metal frame that is placed over a 1.0 ft. x 1.0 ft. x 4 in. deep stainless steel bottomless box frame that
is buried to its rim in the stream substrate. The opening of the net is placed upstream. All substrate
within the rim, and down to the bottom of the box frame, are agitated (or removed) to dislodge
organisms, which drift into the collection bag (Rabeni, 1996).
The standard mesh size of the net is 9 threads/cm. While a smaller mesh size might increase the
number of organism collected, it also will clog more easily and exert more resistance to the current
than a larger mesh. This sampler is specific for the macrobenthos; many microcomponents of the
benthos are not collected (APHA, 1985).
B. The Ponar Grab Sampler
This sampler is used extensively in medium to deep rivers, lakes and reservoirs. It has side plates and
a screen on top of the sample compartment to prevent sample loss during closure. With one set of
weights, the standard 23 X 23 cm sampler weighs 20 kg. The large surface disturbance associated
with the Ponar grab can be reduced by installing hinged, rather than fixed, screen tops, thereby
reducing the pressure wave associated with the samplers descent. This sampler is best used for sand,
gravel, or small rocks with mud but can be used in all substrates except bedrock (APHA, 1985).
C. Multiple-Plate or Modified Hester-Dendy Sampler
The multiple-plate sample is constructed of 0.3 cm thick-tempered hardboard with 7.6 cm round
plates and 2.5 cm round spacers that have center-drilled 1.6 cm holes. The plates and spacers are
held together by a 0.63 cm diameter eye bolt and a nut at the bottom. There are 14 large plates and
24 spaces in each sampler. With the modified Hester-Dendy sampler, the top nine plates are
separated by a single spacer, and plate 10 by two spacers. Plates 11 and 12 are separated by three
spacers, and Plates 13 and 14 by four spacers. The sampler is approximately 14 cm long and 7.6 cm
2
50 in diameter, and has an exposed surface area of approximately 1160 cm and weighs about 0.45 kg.
Macroinvertebrate
Because it is cylindrical, the sampler fits a wide mouth container for shipping and storage (APHA,
1985). This sampler can be hung by a line into a lake or stream. It can be examined in total for
colonization by macroinvertebrates, or the plates can be removed and examined (both qualitatively
and quantitatively) one or more at a time over a period of days or weeks.
PROCEDURE
Equipment and supplies:
1. Surber Sampling
•
•
•
•
•
•
•
•
•
•
•
•
Surber sampler
6 glass vials with caps
3 white porcelain cake pans (with grids)
9 strips of Rite-in-the-Rain labels
3 artist paint brushes
1 liter of 10% formalin solution
field keys to macroinvertebrates
field notebooks and field data sheets
3 clip boards w/rain cover
hip boots or waders
polarized sun glasses (optional)
raingear (if needed)
2. Ponar Sampling
•
•
•
•
•
•
•
•
•
•
Ponar grab sampler and 50 ft. line
Stainless steel sieve tray
3 glass vials and caps
1 five gallon bucket
3 white porcelain cake pans (with grids)
3 strips of Rite-in-the-Rain labels
3 artist paint brushes
1 liter of 10% formalin solution
field keys to macroinvertebrates
field notebooks and data sheets
3. Multiple-Plate Sampling
•
•
•
•
3 Hester Dendy multiple-plate samplers
1 coil of nylon cord to attach sampler
knife
field notebook and data sheets
Teams of 2-3 students per team will practice using the Surber sampler in _________Creek.
51
Macroinvertebrate
LAB PRODUCTS
A. Data sheets for the Surber and Ponar portions of this lab will be turned in to the instructor.
B. Data sheets for the Hester Dendy sampler will be due at the end of the quarter when you turn
in your field notebooks for final grading.
C. Prepare a team write-up (1-2 pp) for this lab exercise. Address the effectiveness of each
method; when?, where?, why?, and how? each is used and the value of macroinvertebrate data
to an overall watershed ecosystem study.
52
Macroinvertebrate
D ate /Time
Common Name
Ge nus/Spe c ie s
#Found
Sk e tc h
Comme nts
Page ___ of ___NAMES: _______________________________________________________________________________
SURBER SAMPLER
DATA SHEET
Location #1 ___________________
Location #2 ___________________
Location #3 ___________________
Replicate # _____
53
Macroinvertebrate
D ate /Time
Location
Location
Location
Replicate
Common N ame
Ge nus/Spe c ie s
#Found
Sk e tc h
Comme nts
NAMES: _____________________________________________________________________________
PONAR GRAB SAMPLER
DATA SHEET
#1 ___________________
#2 ___________________
#3 ___________________
# _____
Page ___ of ___
54
Macroinvertebrate
D ate /Time
Location
Location
Location
Replicate
Common N ame
Ge nus/Spe c ie s
#Found
Sk e tc h
Comme nts
NAMES: _____________________________________________________________________________
HESTER-DENDY
MULTIPLE-PLATE SAMPLER
DATA SHEET
#1 ___________________
#2 ___________________
#3 ___________________
# _____
Page ___ of ___
55
Macroinvertebrate
NOTES FOR INSTRUCTORS
Explain the following prior to or as students do the lab.
SURBER SAMPLER
Teams of 2-3 students per team should practice using the Surber sampler in _________Creek.
Duplicate Surber samples should be collected at three stations along your assigned stream reach.
Sample at the beginning, middle and up-current end of your reach. Place the contents of your
Surber bag into one white porcelain cake pan (with grid). Sort organisms into different sections of
the pan using the artists paint brushes provided. Key out your specimens by common name,
phylum, class, and order, if possible. Some specimens may be identified to the family, genus and
species level. Specimens may be keyed at a later time too. Count the numbers of each species
found. After sorting, keying out and enumerating your macroinvertebrates, place the contents in a
glass vial, label with location, date/time, type of sampler used, replicate number, name of
collector, and other pertinent information using the Rite-in-the-Rain strips. Use a pencil to record
your information. Record observations in your field notebooks and data on your field data sheets.
PONAR GRAB SAMPLER
In your teams of 2-3, take three (3) samples with the Ponar grab from different locations on the
lake. Pour your samples into the stainless steel sieve tray. Rinse with lake water from your 5-gallon
bucket until all of the mud is washed away. Remove all foreign objects (rocks, wood chips, leaves
etc.) by hand. Rinse your organisms into your white porcelain pan. Sort according to like organisms. Key them out to the greatest detail possible. Count each species. Make entries in your field
notebooks and on your data sheets. After sorting, keying, counting and recording, place the
contents of your tray into your glass vials. Label with the same information you used for your
Surber samples.
HESTER-DENDY (HD) MULTIPLE-PLATE SAMPLER
Your team will place one HD multiple-plate sampler in each of three (3) locations. Your multipleplate sampler work is done for today. You will be responsible for removing two (2) discs each week
(beginning next week) until the quarter ends (or you run out of discs). Counting will include
scraping your disc into a white porcelain pan and keying out and counting as in the Surber and
Ponar protocols. You should also examine the phytoplankton and zooplankton. Calculate the
surface area of the discs so that you can quantify the density of each species of macro-invertebrates
and the phyto- and zooplankton. Fill out your data sheets and field notebooks each time you collect
and analyze your samples. Your data will be used for your final research report.
56
Macroinvertebrate
•FOR INSTRUCTORS•
REFERENCES
American Public Health Association, 1985. Standard Methods for the Examination of Water and
Wastewater. 16th edition. Washington, D.C., 1268 pp.
Burch, J.B. 1982. Freshwater snails (Mollusca: Gastropoda) of North America. EPA-600-82026. U.S. Environmental Protection Agency.
Clarke, A.H. 1981. The freshwater Mollusca of Canada. National Museum of Natural Science/
National Museum Canada.
Hobbs, H.H., Fr. 1972. Crayfish (Astacidae) of North and Middle America. Biota of freshwater
ecosystems. Ident. Manual No. 9, U.S. Environmental Protection Agency, U.S. Government
Printing Off., Washington D.C.
Holsinger, J.R. 1972. The freshwater amphipod crustaceans (Gammanidae) of North America.
Biota of fresh water ecosystems. Ident. Manual No. 5, U.S. Environmental Protection Agency, U.S.
Government Printing Off., Washington, D.C.
McCafferty, W.P. 1981. Aquatic entomology. The fisherman’s and ecologist’s illustrated guide to
insects and their relatives. Science Books International, Boston, Mass.
Merritt, R.W. and K.W. Cummings, eds. 1983. An introduction to the aquatic insects of North
America, 2nd ed. Kendall/Hunt Publishing Co., Dubuque, Iowa.
Murphy, B.R. and D.W. Willis, editors, 1996. Fisheries Techniques, 2nd edition. American Fisheries
Society. Bethesda, Maryland.
Rabeni, C.F., B.R. Murphy and D.W. Willis, editors. 1996. Fisheries Techniques. 2nd edition.
American Fisheries Society. Pages 335-352.
•FOR INSTRUCTORS•
57
Macroinvertebrate
FISH 221
Mark-Recapture Method
Mark-Recapture Method
(Lincoln Index)
INTRODUCTION
The mark-recapture method of estimating animal density is used frequently in fisheries and wildlife
studies (Pollack, et. al., 1990). This method was introduced in the late nineteenth century by C.G.
Peterson and later re-introduced for studying bird populations by F.C. Lincoln in late 1930 (Begon,
1980). The method is based upon the fact that if we capture and mark some members of a total
population, recapture them and then sample the population again to find out what proportion of
the sample bears the mark, we can then, by proportionality, estimate the total population size.
The proportionality is:
Rearranging this proportion we get the more familiar formula:
l = Mn
Estimated population N
r
where:
l= estimated total size of the population
N
M = number of animals marked in the population
n = total animals caught in the recapture sample
r = number marked in the recapture sample
NOTE: This estimate is for the date of marking, not the date of recapture.
This method is commonly used because it is relatively easy to accomplish, relatively inexpensive
and can give fair estimates of population density in a rather short time frame. It is, however,
somewhat inaccurate because:
l is not a very precise estimator of N unless the sample size is large, or a
1. N
high proportion of the population is marked.
58
MarkRecapture
2. Several assumptions may or may not be valid in an actual population
(Cormack, 1993). These assumptions are:
a) “marks” are not lost
b) there is no recruitment into the population by reproduction, growth or immigra
tion, or emigration out of the population
c) marked and unmarked animals behave alike (mortality rates, activity, and
response to traps are the same)
Jolly (1965) devised a capture-recapture statistical model that considers death and immigrating
which gives more validity to this method.
PROCEDURE
Equipment and Supplies:
• 1 bag each of white and brown beans (of roughly the same size)
• 1 plastic bucket
• 5 hand calculators (with statistical functions)
• release-recapture data sheet
In this exercise we will use a small plastic bucket (represents a lake) filled with white beans
(juvenile coho salmon). The object is to estimate the coho population density in the lake.
The instructor will begin by removing a handful of white beans from the bucket. These will be
counted and put aside. An equal number of brown beans (representing those coho that were
captured and marked) are put back into the bucket. All beans are thoroughly mixed.
Each team of students will then use a plastic cup (with a capacity of about 75-100 beans) to
simulate a beach seine and scoop up a recaptured sample. Beans should be thoroughly mixed
between scoops (seines). Students will enter their data in the space below. The instructor will
l. Also
supply the figure for the number of marked beans (coho) in the population. Compute N
enter other teams’ estimates as they become available.
Number marked in sample (r) = __________
Sample size (n) = __________
Total marked in population (M) = __________
l) = __________
Your estimate of total population size (N
l ) = __________ , __________ , __________
Other team estimates of total population size (N
59
MarkRecapture
DISCUSSION QUESTIONS
l from your instructor. How far off was the best estimate derived by
1. Get the actual value for N
sampling? The worst estimate? Calculate the mean (x ) of all the estimates. Calculate the median
of all the estimates; and the mode, range, and standard deviation. Was this sampling method a
close approximation of the actual population?
2. Why is the Lincoln index estimate for the time of marking rather than the time of recapture?
3. List several methods for capturing fish, amphibians, reptiles, birds and mammals.
60
MarkRecapture
4. List several methods for marking fish, amphibians, reptiles, birds and mammals.
5. Considering the actual population of coho in a lake, try to assess which of the assumptions
necessary to validate the mark-recapture method—consider how it might be violated, and specifically, how.
6. What are “trap-shy” and “trap-prone” animals? How might each of them affect your estimate of
population size?
61
MarkRecapture
7. Why isn’t mortality considered an assumption for the mark-recapture method whereas reproduction is?
l, would be too high or too low in each of the following
8. Indicate whether your estimate, N
circumstances:
a. Some coho lose their marks.
b. Immigration occurs.
c. Marked coho have higher mortality rates.
d. Marked coho are more likely to be seined.
e. Marked coho are more sluggish than unmarked ones.
62
MarkRecapture
Confidence Intervals:
It is desirable when using the Lincoln index to calculate a confidence interval, the interval within
which you are confident (with a specific probability) that the true population lies. In general, the
value we add to and subtract from (for the upper and lower confidence limits) any estimate is
given byK S 2, where S 2 is a variance and K is the normal curve variate. Unfortunately, the
variance is not straight forward for the Lincoln index population estimate. Therefore, the formula
we will use for our exercise today is:
S 2 = M 2 n( n − r )
r3
Enter here your estimate of S 2 : __________
Take the square root of S 2 and enter here your value of
S 2 : ____________
The useful normal curve variates (K) for the 95% probability is 1.96 (taken from a table of normal
curve variates at different probability levels). Thus, now multiply 1.96 times S and enter it here:
__________
l) to give the upper and
This is the value you will add to and subtract from your estimate of N (N
lower limits of the confidence interval.
Give those limits here: __________
Now write a comprehensive statement in the space below that gives the confidence intervals for
the estimate of the coho population in the lake.
_____________________________________________________________________________________________________________________________________________________________________________________________________________________________________________________________
_________________________________________________________________________________________________________________________________________
To get the 99% confidence limits for our population estimate, multiply S times 2.58 (the normal
variate for 99% probability), and then add and subtract this value from your estimate of N.
Give those limits here: __________
[Some of the formuli listed above were taken from: Cox, G.W. 1996. Laboratory Manual of
General Ecology. Seventh Edition]
63
MarkRecapture
LAB PRODUCTS
During the lab:
•
•
•
•
Following the Lab:
•
•
Carry out the bean-counting and calculations.
Participate in the discussion.
Complete your calculations and fill in your data sheets and calculations.
l) for the
Exchange data with other teams. Use your estimate of N (N
calculations of confidence intervals (CI).
Clean up your lab space. Separate brown and white beans and return them to
their appropriate containers.
Turn in one team data sheet with names of all team members in the upper
right hand corner of the first page.
Turn in a 2-3 page team report that addresses the questions in the lab exercise. Be as specific as
possible!
REFERENCES
Begon, M. 1980. Investigating Animal Abundance: Capture-Recapture for Biologists. University
Press, Baltimore, Maryland.
Cormack, R.M. 1993. Variances of Mark-Recapture Estimates. Biometrika 49-118-1193.
Cox, G.W. 1996. Laboratory Manual of General Ecology, Seventh Edition. Wm.C. Brawn
Publishers, Dubuque, IA. pp. 66-67.
Jolly, K G.M. 1965. Explicit Estimates from Capture-Recapture Data with Both Death and
Immigration-Stochastic Model. Biometrika 52: 225-247.
Pollack, K.H., J.D. Nicols, C. Browne, and J.E. Hines. 1990. Statistical Inferences for CaptureRecapture Experiments. Wildlife Monographs No. 107.
Identification Guides:
Leonard, W.P., H.A. Brown, L.L.C. Jones, K.R. McAllister, and R.M. Storm. 1993. Amphibians
of Washington and Oregon. Seattle Audubon Society, Seattle, WA. ISBN 0-914516-10-8
Udvardy, M.D.F. 1985. The Audubon Society Field Guide to North American birds. Western
Region. Alfred A. Knopf, New York.
64
MarkRecapture
FISH 221
Gas Supersaturation
Gas Supersaturation
INTRODUCTION
A. What Is Supersaturation?
Dissolved gasses are present in all fish-bearing waters, usually entering the water directly from the
air. Normally, the dissolved gas pressure is equal to the atmospheric (barometric) pressure just
above the air-water interface (14.7 PSI or 760 MM of mercury at sea level). NOTE: 100% saturation
is considered normal and ideal.
Occasionally, natural or man-made conditions occur that upset this balance. Supersaturation can be
caused by dam spillways, waterfalls, power plant outflows, algae blooms and other sources. If the %
saturation goes higher than 110%, aquatic life is threatened and fish kills can occur. If the %
saturation goes below 80%, not enough dissolved oxygen is present to support aquatic life.
Undersaturation (de-aerated waters, below 100% saturation) can occur in some well or spring waters
and in still or stagnant ponds.
Supersaturation causes a disease in fish called the “gas-bubble disease.” Air bubbles come out of
solution in the blood vessels of fish causing blockage, rupture and death. This disease is similar to
the bends experienced by human divers coming to the surface too quickly from a deep dive.
B. How was the Saturometer invented?
Large fish kills in the early 1970’s on the Columbia and Snake river systems in the Pacific
Northwest prompted the E.P.A. to take action to solve the problems. An inexpensive, field
portable, accurate instrument to measure dissolved gas pressures at many various locations was
needed. Dr. Ray F. Weiss, with the Scripps Institute of Oceanography, in conjunction with people
from Region 10, E.P.A. in Seattle, designed the first operational saturometer. ECO Enterprises
redesigned and started manufacturing these instruments in 1972.
Your instructor will provide you with pertinent background information, standards, and protocols.
65
Gas
Supersaturation
PROCEDURE
We will visit our local fish hatchery. Students will work in teams of 2-4
1. Following the procedure demonstrated by the instructor or assistant, determine the barometric pressure using a mercurial barometer. This reading may be verified by checking with a
local airport.
2. Using the ES-2 Weiss Saturometer (the long handled type) and following the correct protocol,
determine the percent (%) saturation for the lake at the floating net pen. Use the same
saturometer to find percent (%) saturation in the creek at the outfall below the lake.
Record your measurements on the field data sheet provided. Exchange your instrument with
other teams when completed.
3. Using the ES-3 Weiss Saturometer (the short-handled meter) determine the percent (%)
saturation in the headbox inside the hatchery. Measure the percent (%) saturation in at least
one (1) of the fish troughs containing coho salmon fry. Record your measurements on the field
data sheet provided. Exchange your instrument with other teams when completed.
4. With your instructor’s direction, perform your calculations, and record them on the data sheets
provided.
EQUIPMENT AND SUPPLIES
•
•
•
•
•
•
field notebook
lab exercise and data sheets
mercurial barometer
ES-2 and ES-3 Weiss saturometer
telephone
hand-held calculator
LAB PRODUCTS
At the end of this lab you will be responsible for:
•
Rinsing both the ES-2 and ES-3 saturometers and stowing them away according to the
hatchery manager’s directions.
•
Complete your data sheets. Remember to obtain data from other teams to calculate mean
and standard deviation for percent (%) saturation measurements taken by your team.
•
Turn in one data sheet for your team at the beginning of the next lab/field period. Remember
that each individual is responsible for the procedure, data collection and calculations for this
exercise.
66
Gas
Supersaturation
67
Gas
Supersaturation
Loc ation 4
Loc ation 3
Loc ation 2
Loc ation 1
Loc ation
Instr ume nt
Use d
Loc ation 2
Te ams S.D . of %
Saturation
Loc ation 3
P atm + D P X 100
P atm
Te ams Ave rag e %
Saturation
Loc ation 4
Te am Me mbe r N ame s
Comments:__________________________________________________________________________________________________
___________________________________________________________________________________________________________
___________________________________________________________________________________________________________
Barometric Pressure (from mercurial barometer) = _____ mm Hg
Barometric Pressure (from airport) = _____ mm Hg
Loc ation 1
=
Pe rc e nt (%)
Saturation
Calculations: Percent (%) Saturation
D ate /Time
GAS SATURATION DATA SHEET
NOTE FOR INSTRUCTORS
NOTE Introductory information in this laboratory exercise has been used with permission from
the “Field Manual for the ES-2 and ES-3 Weiss Saturometers,” ECO Enterprises, 2821 N.E. 55th,
Seattle, WA 98105. Be sure to check the sites where sampling is undertaken in this exercise to
ensure you have access to similar facilities for your students.
The following is general information to share with your students. Important protocols for
conducting this lab are included.
A. Principles of Operation
The saturometer operates basically as an artificial fish gill. Dimethyl silicone rubber tubing is used
for the sensing membrane. This small, hollow tubing will allow the passage of dissolved gasses from
the outside (water) to the interior (pressure gauge, referenced to local atmospheric pressure) for
readout. Over 300 square inches (200 ft.) of this tubing is used in the ES-2 instrument.
The tubing (.025” O.D. x .012” I.D.) is wound on frames into a compact sensor. A perforated outer
shell protects the membrane tubing from damage and allows rapid water transfer. A manually
operated water-jet pump emits high velocity streams of water across the tubing, accelerating water
transfer and removing air bubbles from the tubing that interfere with dissolved gas diffusion. The
water-jet pump is not needed in undersaturated waters as no bubbles can form.
A specially calibrated low-volume pressure gauge is used to measure the dissolved gas pressure. This
gauge measures from -100 to +400 mm of mercury with a full-scale accuracy of better than ± 1%.
A pressure relief valve is located just behind the gauge to quickly vent internal pressure after a
reading is taken, for testing, leak detection and calibration purposes.
B. Saturometer Use in Rivers, Lakes & Streams
Before industrial man started utilizing our waters for power generation, irrigation and flood control,
the only source of supersaturation was waterfalls—which trapped atmospheric gasses and carried
them into deep pools where the gasses went into solution. Nature’s way of dissipating the resulting
supersaturated condition was to take these waters over turbulent, shallow sections of the river that
agitated the water, bringing the dissolved gasses in contact with the air-water interface where they
could dissipate and return to the atmosphere.
The advent of large dams on many fish-bearing rivers and streams eliminated these turbulent
sections by creating large, deep reservoirs that prevented effective dissipation of excess dissolved gas
pressures. Dam spillways, when in operation and not properly designed, contributed more to the
supersaturation levels. Hot water from power generating facilities (nuclear or other) entering these
waters further contributed to this problem.
Unlike the air we breathe, the dissolved gas pressure and composition in water are highly variable
and can be adversely affected by many different factors and conditions. The location, control,
68
Gas
Supersaturation
•FOR INSTRUCTORS•
regulation or elimination of these undesirable factors falls under the jurisdiction of many diverse
disciplines: regulatory agencies such as the E.P.A., state, federal and private fisheries agencies,
power companies and utility districts, the Departments of the Interior and Commerce, and private
industry are all involved in controlling and maintaining a healthy aquatic environment for fish and
fisheries-based industries.
The Weiss Saturometer, as a field portable accurate instrument to monitor dissolved gas parameters,
has proven to be a valuable asset on a world-wide basis. Both rugged and inexpensive, fast, accurate
readings can be obtained. At the monitoring site, the sensor membrane section is immersed in the
water and the valve closed. A few minutes of agitation with the water pump (swishing back and
forth) will indicate whether the water is supersaturated (pointer on gauge will move clockwise, +) or
undersaturated (pointer on gauge will move counterclockwise, – ). If undersaturated water is
evident, no agitation or pumping is needed as air bubbles will not form on the tubing. If supersaturated conditions are present, vigorous agitation and pumping every 30 seconds of agitation should
be used. If no change in the reading occurs, equilibration has been reached and that reading should
be noted.
Other information should be recorded, preferably on a data sheet, and should include date, time,
location, barometric pressure (either measured or calculated from weather service data and altitude
charts), water and air temperature. If % nitrogen is needed, a dissolved oxygen determination
should be made with either a D.O. probe or the Winkler method.
If tests at difficult sites or from a boat are required, it may be wise to use the wrist-straps on the
ES-2 or tie a line around the handle to prevent loss of either type of instrument.
An alternative method of obtaining a saturometer reading is preferred by some—a pump or syringe
is used to apply a positive pressure (or by sucking on the valve opening to provide a vacuum), then
the valve is shut so a reading 50 to 100 mmHg above the expected reading is obtained. The
instrument, immersed in the water to be tested, is then allowed to bleed down to the equilibration
point. Agitation may still be needed in supersaturated waters. Both methods can be employed and
the average value computed for very accurate readings. Either method takes generally 4-15
minutes to obtain a reading.
Factors that can alter the equilibration time and accuracy of the readings include:
1.
Condition of the tubing—periodic cleansing of the membrane tubing in a detergent
solution is recommended.
2.
Temperature of the tubing—hot water immersion just prior to taking a reading speeds up
reading time.
3.
Water conditions—oil, industrial waste, other pollutants can coat the tubing adversely
affecting equilibration time and instrument accuracy.
4.
Restrictions to gas flow—swagelok fittings being tightened too far can pinch off the teflon
tubing used to transfer pressure to the gauge. There are four “frame” windings in each
•FOR INSTRUCTORS•
69
Gas
Supersaturation
instrument and eight membrane tubing “ends” brought into the accumulator manifold. A
pinch-off in one of the membrane frames will generally not impede accurate readings as
the gas has essentially two directions it can flow to get to the manifold and gauge.
5.
Leaks—a leak anywhere, no matter how small, will adversely affect both the equilibration
time and final reading. NOTE: The instrument must be maintained as a sealed system!
Equilibration is defined as the point where the internal (dissolved) gas pressure equals the
atmospheric pressure at the measurement location plus the gauge diaphragm and spring pressure
that moves the gauge pointer to its stable position. After readings are taken, the lower section of
the instrument should be rinsed off with fresh water, dried and the instrument should be stored in
a plastic bag or suitable container away from excess humidity, direct sunlight or heat extremes.
If slow reading times are experienced, it may be due to the presence of water in the tubing. Longterm immersion is not recommended as some water vapor can traverse the membrane tubing wall
and condense inside the tubing, causing a blockage condition. If this condition is expected, the
instrument can be dried out in an electric oven at 180o F for one hour.
CAUTION!! Do not dry the instrument or expose to open flame or fire as the tubing may be
destroyed. Care should be taken to keep the gauge dry and free from corrosion. In taking field
readings, do not allow foreign objects such as sticks, sand, etc., to enter the sensor area.
C. Saturometer Use In Hatcheries, Research Facilities
An active saturometer-monitoring program in government or private hatcheries can prove of great
value in preventing fish kills due to supersaturation or oxygen depletion and in regulating the
aquatic atmosphere through dynamic control of aerators and source water selection, metering and
composition control. Many hatcheries now employ almost totally artificial (man-made) control of
the amount and type of dissolved gases present in their fish bearing waters. The advent of intensive fish farming programs and modern research facilities intensifies the necessity to accurately
measure the composition and amount of dissolved gasses present. It has been proven that fish
kills due to dissolved gas supersaturation are not necessarily due to nitrogen alone, but the sum
total of all dissolved gas partial pressures—the exact condition that the Weiss Saturometers
measure!
REMINDER: It is still a necessity to maintain a proper nitrogen/oxygen balance to assure the fish
have enough oxygen present to survive!
Many hatcheries, fish farm facilities, and research institutes are now altering this nitrogen/oxygen
balance to obtain optimum growth rates of fingerlings and fry, often with great success. The
necessity of maintaining these de-aerated and then oxygen enriched waters at the optimal %
saturation pressure dictates an active monitor and control program due to the complexity of these
systems. One failure or mistake gone unnoticed for a given length of time may cause the loss of
millions of young fish.
70
Gas
Supersaturation
•FOR INSTRUCTORS•
A sudden change in source water composition and % saturation occasionally occurs. The first
warning signs of trouble without a saturometer-monitoring program are usually a lot of belly-up
fish. With a monitoring program, such conditions can be detected and corrected before fatalities
occur. The same is true if the aerator and circulations pump systems should a leak (usually at a
fitting) in a high pressure or high velocity section of piping, or in a suction section on the inlet side
of a pump occur, where atmospheric air can be injected into that point via venturi action causing
supersaturation to be present past that point. If the condition persists, fatalities can occur. With a
spot check saturometer-monitoring program, the leak can be located and fixed before fatal conditions occur.
Stagnant conditions can produce algae blooms, often in larger holding ponds. They produce
prodigious amounts of dissolved oxygen that can reach harmful or fatal levels. By taking readings
on a grid pattern using triangulation measurements, a thermal map can be constructed, pinpointing
the algae bloom source. Localized herbicide then can be selectively applied to control this condition
without poisoning the entire pond. And the presence of too many fish in a given body of water can
lead to oxygen depletion. These conditions can be detected and corrected with an active
saturometer monitoring program before fatalities develop.
These basic operating procedures outlined are used in hatcheries and other facilities. The ES-3
“mini” or lab saturometer may be better suited for use in smaller size fish tanks as it occupies less
space and removes less dissolved gas from the water than the larger ES-2 instrument. The longer
handle on the ES-2, however, allows hatchery personnel better access to outdoor holding ponds
which may have water levels a few feet below the ground surface.
Saturometers can be sterilized with acetone, boiling water, dilute chlorine or other bacterial agents to
prevent the possible spread of diseases from tank to tank or pond to pond.
D. Calculating Percent Saturation
Before immersion of the saturometer sensor section in water, the pressure gauge will indicate a zero
reading. This is the point at which the pressure inside the instrument is equal to the local atmospheric (barometric) pressure. After a saturometer reading is taken, the gauge pointer will indicate a
positive or negative dissolved gas pressure reading in millimeters of mercury. To convert this
number into % saturation, the atmospheric pressure should be measured with an aneroid or
mercurial barometer or calculated from altitude charts and local weather bureau or airport data. The
following formula is then used:
(Equation #1) Percent (%) Sat. = P atm + D P X 100
P atm
•FOR INSTRUCTORS•
71
Gas
Supersaturation
Where % Sat. is the total dissolved gas saturation (supersaturation or undersaturation), P atm =
the local barometric pressure in mm Hg (760 mm Hg nominal at sea level), D P is the saturometer
reading in mm Hg.
Example:
Barometric pressure is 765 mm Hg, P is + 100 mm Hg, therefore:
The actual dissolved gas pressure present in the water is 865 mm Hg.
NOTE: For most applications this number (% Sat.) is the only number needed to assure safety to
the aquatic environment and to assure legal requirements are within proper limits.
E. Conversion Factors
Atomic Values:
O2 = 22.392 liters/mole
N2 = 22.403 liters/mole
Ar = 22.390 liters/mole
Atomic Weights:
O2= 31.9988 grams/mole
N2 = 28.0134 grams/mole
Ar = 39.948 grams/mole
Composition of air:
O2 = 20.946%, N2 = 78.084%, Ar = .934%
Concentration:
1 ppk = 1 mg/liter (approx./fresh water)
.001 gm/L = 1 oz./7400 gallons (approx./fresh water)
1 gm/liter = 1000 parts per million (approx./fresh water)
Conversion of ml/L to mg/L:
1 ml/L O2 = 1.42903 mg/L
1 ml/L N2 = 1.25043 mg/L
1 ml/L Ar = 1.78419 mg/L
Distance:
1 ft. = .3048 meters
1 meter = 3.281 feet
Hydrostatic Pressure:
1 meter of fresh water @ 20o C = 73.43 mm Hg
1 foot of fresh water @ 20o C = 22.89 mm Hg
1 meter of seawater @ 35 ppt, 20o C = 74.88 mm Hg
1 foot of seawater @ 35 ppt, 20o C = 23.34 mm Hg
72
Gas
Supersaturation
Pressure: 1 Std. atmosphere = 760 mm Hg = 1.013 Bar = 10.35 meter (32.2 ft.) of fresh water at
20o C = 10.15 meter (32.56 ft.) of seawater at 35 ppt and 20o C = 14.22 lbs./sq. in. = 101.325
kilopascals = 760 torr @ 0o C = 1 kg/cm2
Temperature: oC = 5/9 (Fo - 32)
F = (9/5 oC) + 32
o
Volume: 1 cc = 6.1 X 10-3 cu. in.
o
K = oC + 273.15
1 cu. ft. = 28.32 liters
REFERENCE
ECO Enterprises, 1985. Weiss saturometer field manual for the ES-2 & ES-3 instruments. ECO Enterprises, 2821 N.E. 55th, Seattle, WA 98105. (206) 525-4784 or (206) 523-9300; also, 1-800-426-6937
(Exc. WA).
73
Gas
Supersaturation
FISH 221
Beach Seining & Otter Trawling
Beach Seining and Otter Trawling
INTRODUCTION
A. Otter Trawls
Otter trawls are commonly used in surveys of demersal species, particularly in marine ecosystems
and estuaries (Murphy and Willis, 1996). The general purpose of otter trawling is to provide indices
of abundance of different species of finfish and shellfish and provide biological information for
understanding their population dynamics. For research purposes, otter trawling is usually conducted
from 5 meters to 150 meters in depth. Many bottom trawl surveys are combined with midwater
trawling and hydroacoustic surveys (Hayes et al., 1996).
One reason for the popularity of trawls is that they sample a discrete area of the bottom, or volume
of the water column, over a specified time. While trawls are quite versatile in the variety of habitats
they can sample, they do have some limitations. Trawls cannot be effectively used on bottom
substrates that can snag the net. Coral reefs, aquatic macrophyte (kelp) beds, and rocky
outcroppings are examples of habitats that can’t be sampled with otter trawls.
B. Beach Seines
A beach seine is a type of encircling net. Encircling nets are used to trap fish actively by surrounding
them in a fence-like wall of netting. Surrounding nets provide several advantages over otter trawls
because the gear is easy to deploy, sampling is rapid, a large area can be sampled, the limits of the
sampling area are well defined, and fish are captured live with minimal trauma (Murphy and Willis,
1996). These nets also allow for live release (Pierce et al., 1990). Beach seines, unlike otter trawls,
can be used without a boat, and can be fished by a single person.
Beach seines are often fished in a semicircle around the targeted fish and dragged to shore, herding
the fish into the net (Murphy and Willis, 1996). Setting and hauling beach seines can be done in
several ways. One method is for a single vessel or person wading in shallow water to fix a towline on
shore and then to set a wing and the bunt offshore before turning in to shore and setting the second
wing and towline. It’s common to set the net up-current or up-wind in some cases, to give the net a
chance to open to its greatest width. However, the lead-line must be well anchored to prevent the
seine from lifting off the bottom if this technique is used. The net is hauled in by dragging in the
towlines until the bunt reaches shore and the catch can be retrieved.
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PROCEDURE
Many studies in which beach seines are used rely on a single haul to represent the fish community in
the area being seined (Allen et al., 1992). These researchers showed that species richness, species
rank, and size distribution of dominant taxa were well represented by the first haul.
During the next two field exercises we will gather data that will either support or refute the above
research findings.
Equipment and Supplies:
•
•
•
•
•
•
•
•
•
•
•
•
•
2 Boston Whaler boats
1 research otter trawl
1 straight beach seine
field notebooks
layered clothing
raingear, hip boots, knee boots and/or waders
this lab exercise and data sheets
6 five gallon plastic buckets
6 plastic specimen tubs
camera and binoculars (optional)
species identification keys (adults and juveniles)
2 YSI dissolved oxygen meters
2 YSI salinity, conductivity, temperature meters
The class will split into two teams (blue and gold crews). The blue team will do beach seining, with
the instructor while the gold team will perform otter trawling with our instructional assistant.
During the next lab/field period, teams will switch; the blue crew doing the otter trawling and the
gold crew beach seining.
A. Beach Seining
Three side-by-side (replicate) beach seines will be made each sampling day. Teams should mark off
three 150’ sections along the beach by placing stakes in the sand/mud. The instructor will provide
on-site details as to which beach seining method to use. Use the data sheets provided to record your
observations and measurements.
You will be recording species of plants and animals found, numbers of each species, and you will be
making some basic water quality measurements (dissolved oxygen, salinity, conductivity and water
temperature) for each replicate.
NOTE: Be sure to record routine observations in your field notebooks!
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B. Otter Trawling
Three (3) replicate otter trawls will be performed. Each trawl will be exactly 5 minutes long (once
the trawl reaches the bottom).
The instructor will provide specific details as to the direction of tows, length of tows, and the
setting and retrieval of the trawl. Like seining, you will be recording species of plants and animals
found, numbers of each species found, and you will be making some basic water quality measurements (dissolved oxygen, salinity, conductivity, and water temperature) for each replicate. And again,
be sure to record routine observations in your field notebooks.
C. Departure and Return
We will leave _________by vans at __________. We will return at __________. Be sure to
develop a check off sheet for this exercise.
NOTE: The instructor will ask you to review your list before we leave on this field exercise.
LAB PRODUCTS
At the end of these lab/field exercises, you will be responsible for:
1. Rinsing the beach seine and trawl net with fresh water, and stretching them out to dry in the
designated area.
2. Rinsing or wiping off all other equipment, boats and gear with fresh water, and temporarily
placing all gear and equipment in the designated drying area. Once it’s dry, stow it where you got
it.
3. Complete your otter trawl and beach seine data summary sheets; make sure your field notebooks
are complete; write a 4-5 page, word processed (or typed) individual research report about your
results for these two lab/field exercises. Reports should contain a title page, table of contents,
introduction, methods and materials, results and discussion, summary and/or conclusions, and
bibliography. Use tables and graphs as needed. Your raw data tables and summaries can go in
your appendix. Photos can be used as figures in your report.
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Otter
REFERENCES
Allen, D.M., S.K. Service, and M.V. Ogburn-Mattews. 1992. Factors influencing the collection efficiency of
estuarine fish. Transactions of the American Fisheries Society 121: 234-244.
APHA 1985. Standard methods for the examination of water and wastewater. 10th edition. American Public
Health Association, Washington D.C. p 1132.
Byrne, C.J., and J.R. Nicolas. 1989. Bottom Trawl Survey Manual. National Marine Fisheries Science
Center, Woods Hole, Massachusetts.
Hayes, D.B., C. Paola Ferrei, and W.W. Taylor. 1996. Active Fish Capture Methods. (pp 193-220) B.R.
Murphy and D.W. Willis, editors. Fisheries Techniques, 2nd edition. American Fisheries Society,
Bethesda, Maryland. 732 pp.
Murphy, B.R. and D.W. Willis, editors. 1996. Fisheries techniques, 2nd edition. American Fisheries
Society, Bethesda, Maryland. 732 pp.
Canadian Journal of Fisheries and Aquatic Sciences. 47: 1004-1010.
Keys to freshwater and marine fishes:
Hart, J.L. 1973. Pacific fishes of Canada. Bulletin 180. Canadian Government Publishing Centre,
Ottawa, Canada. ISBN 0-660-10459-8.
Lee, D.S., C.R. Gilbert, C.H. Hocutt, R.E. Jenkins, D/E. McCallister, and J.K. Stauffer, Fr. 1980.
Atlas of North American freshwater fishes. Publ. No. 1980-12, North Carolina Biological Survey.
McClane, A.J. 1974. Freshwater fishes of North America. Holt, Reibart and Winston, New York, New
York.
Marrow, J.E. 1980. The freshwater fishes of Alaska. Alaska Northwest Publishing Co., Anchorage,
AK.
Scott, W.B. and E.J. Crossman. 1973. Bulletin 184. Freshwater fishes of Canada. Canadian Government Publishing Centre, Ottawa, Canada. ISBN 0-660-10239-0.
77
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Otter
Common N ame
Conductivity: ____________________ µmhos
# S a m pl e d
Comme nts
Water Temperature: ____________________ oC
Salinity: ____________________ o/o (ppt)
Ge nus & Spe c ie s
Beach Seine Data Sheet
Beach Seine #: _____
Stage of Tide: ____________________
Dissolved Oxygen: ____________________mg/L
D ate /Time
Names: __________________
Location: ____________________
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Beach
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Otter
Com m on N am e
Ge nus & Spe c ie s
Date of Collection:_________
Location: _________________
Page ___ of ___
S.D . = __________
Se ine #_________
_
x = __________
Se ine #_________
Se ine #_________
Total N um be r Pe r Se ine
Beach Seine Data Summary
Com m e nts
Names: ____________________
____________________
____________________
____________________
Comme nts
Water Temperature: ____________________ oC
# Sample d
Conductivity: _________________________ µmhos
Ge nus & Spe c ie s
Salinity: ____________________ o/o (ppt)
Common N ame
Estimated Length of tow: ____________________ meters
Otter Trawling Data Sheet
Trawl #: _____
Dissolved Oxygen: ____________________mg/L
D ate /Time
Stage of Tide: ____________________
Location: ____________________
Names________________
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81
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Otter
Com m on N am e
Ge nus & Spe c ie s
Date of Collection:_________
Location: _________________
Page ___ of ___
S.D . = __________
Tr awl #_________
_
x = __________
Tr awl #_________
Tr awl #_________
Total N um be r Pe r Se ine
Otter Trawl Data Summary
Com m e nts
Names: ____________________
____________________
____________________
____________________
NOTE FOR INSTRUCTORS
NOTE: Other types of sampling such as trap netting or gill netting may be substituted for the otter trawling if
desired, as otter trawling is specific to marine sampling and specific equipment, such as boats with sufficient horsepower, is required for this exercise.
BACKGROUND INFORMATION
A. Otter Trawls
Otter trawls are commonly used in surveys of demersal species, particularly in marine ecosystems
and estuaries (Murphy and Willis, 1996). The general purpose of otter trawling is to provide indices
of abundance of different species of finfish and shellfish and provide biological information for
understanding their population dynamics. For research purposes, otter trawling is usually conducted
from 5m down to 150 m in depth. Many bottom trawl surveys are combined with midwater trawling and hydroacoustic surveys (Hayes, Paola Ferrei, and Taylor, 1996).
A typical otter trawl features a head rope (with floats), a foot rope (with a chain to weigh the net
down and scour the bottom), top and lower wings, a mouth and cod end, bridles that connect to a
pair of trawl doors (that flare outward to keep the wings of the net open when towed), and two
towing ropes that connect to a single or double set of booms on the towing vessel (Byrne and
Nicolas, 1989).
Otter trawls are towed along the bottom. As the net is towed through the water, fish entering the
net eventually tire and are “funneled” to the cod end where they are held until the net is retrieved.
Most trawls are designed so that the cod end is tied shut while fishing and can be opened after
retrieval, to make removing fish from the net easier (Hayes, Paola Ferrei, and Taylor, 1996). Most
bottom trawls for scientific programs use otter doors. Otter doors for bottom trawls are relatively
heavy in order to keep the net on the bottom, as well as to keep the wings of the net spread open
(APHA, 1985). Most otter doors are rectangular and keep lateral pressure on the net by having the
towing warps attached to front brackets, while bridles are attached to the back brackets. The bottoms of the doors typically have metal shoes to protect the doors from abrasion with the substrate
(Byrne and Nicolas, 1989).
One reason for the popularity of trawls is that they sample a discrete area of the bottom, or volume
of the water column, over a specified time. While trawls are quite versatile in the variety of habitats
they can sample, they do have some limitations. Trawls cannot be effectively fished on bottom
substrates that can snag the net. Coral reefs, aquatic macrophyte (kelp) beds, and rocky
outcroppings are examples of habitats that can’t be sampled with otter trawls.
Unless the trawl net is relatively small, it generally requires a powerful vessel to pull the net along the
bottom. It also takes two or more people to handle an otter trawl safely, both in the water and when
the net is on-board.
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•FOR INSTRUCTORS•
B. Beach Seines
A beach seine is a type of encircling net. Encircling nets are used to trap fish actively by surrounding
them in a fence-like wall of netting. Surrounding nets provide several advantages over otter trawls
because the gear is easy to deploy, sampling is rapid, a large area can be sampled, the limits of the
sampling area are well defined, and fish are captured live with minimal trauma (Murphy and Willis,
1996). These nets also allow for live release (Pierce et al, 1990). Beach seines, unlike otter trawls,
can be fished without a boat and can be operated by a single person.
The beach seine is made of mesh of uniform size and consists of two wings and a bunt section that
holds the catch. In some seines, the bunt is enlarged to form a bag in the netting. The wings form a
long vertical wall that funnels fish to the bunt. The beach seine also features a cork line at the top
and lead line at the bottom of the net. A 5-6 foot tall pole is also attached to each wing. Towing
warps or bridles can be attached to these poles.
Beach or haul seines are typically used in shallow waters where the net wall can extend from the
surface to the bottom. Seines with a bagged bunt work better in lakes; bagless seines are used in
rivers.
Beach seines are often fished in a semicircle around the targeted fish and dragged to shore, herding
the fish into the net (Murphy and Willis, 1996). Setting and hauling beach seines can be done in
several ways. One method is for a single vessel or person wading in shallow water to fix a towline on
shore and then to set a wing and the bunt offshore before turning in to shore and setting the second
wing and towline. It’s common to set the net up-current or up-wind in some cases, to give the net a
chance to open to its greatest width. However, the lead-line must be well anchored to prevent the
seine from lifting off the bottom if this technique is used. The net is hauled in by dragging in the
towlines until the bunt reaches shore and the catch can be retrieved.
A second method requires two vessels or two people wading in shallow water. In this case, the two
vessels or people start together at the farthest point from the beach and set the bag. They open the
seine parallel to the beach, each setting a wing and towline. Then the seine is pulled toward the
shore, and the wings are brought together. The bag with the catch is held between the vessels or
hauled ashore where the catch can be collected. Another common method of seining is for two
fishers to set the net perpendicular to shore and then drag the net parallel to shore. At the end of
the tow, the person farthest out turns in to shore and both wings are pulled ashore (Murphy and
Willis, 1996).
•FOR INSTRUCTORS•
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FISH 221
Nekton Sampling
Nekton Sampling
INTRODUCTION
Nekton are all the larger, aquatic, free-swimming animals in lakes, seas, and ponds, whose movements are largely independent of currents and waves. Some species considered to be nekton include
fishes, squids and whales.
PROCEDURE
Today we will perform nekton sampling in Lake____________. We will be using hook and line to
catch and tag resident cutthroat trout. We will use the methods from the Mark and Recapture lab
to mark, release and recapture the cutthroat.
Equipment and Supplies:
• light spinning rod and reel (a hand line will do)
• bobbers
• worms (or other preferred baits)
• #6 or #8 worm hooks
• polarized sun glasses (optional)
• dip net
• spaghetti tags and tagging gun
• field notebook and nekton data sheets
• a current & valid Washington State fishing license
• keys to freshwater fishes (adults and juveniles)
• Washington State Fishing Regulations
• ...and don’t forget to bring your luck!
You are welcome to fish anywhere you like in Lake __________. You can fish individually or in
teams of two.
Identify, count, and release all of the different species you catch today. Tag and release only
cutthroat trout. Record all catch data on the data sheets provided.
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Nekton
LAB PRODUCTS
1. Read over this lab exercise and the current state fishing regulations for Western Washington
lowland lakes. Also review the mark-recapture protocol.
2. Fish like crazy for 3 hours and 50 minutes!!
3. Record your catch data.
4. Turn in a 1-2 page individual report at the beginning of the next lab. Include advantages and
disadvantages of hook and line as a nekton sampling method. Try to describe the best
technique you can for catching cutthroat trout with hook and line (you are encouraged to
consult with some of the more expert fishers in our class for this information!).
Both individual and class results will be summarized. Try to explain the results you got and
how they compared with data for the entire class.
REFERENCES
Eschmeyer, W.N. and E.S. Herald. 1983. A field guide to Pacific coast fishes of North America.
Houghton Mifflin Co., Boston, Mass.
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Nekton
Common N ame
Ge nus and
Spe c ie s
N umbe r
Caug ht
N umbe r
M ar k e d
N umbe r
Re le ase d
General Information (techniques used; bait used; number of fish injured or killed;
etc.):
D ate /Time
Nekton Sampling
Individual Data Sheet
Location (General): _____________________________
Specific Location: ______________________________________________
Samplers: ____________________
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Nekton
Comme nts
87
Nekton
D ate /Time
Common N ame
Ge nus and
Spe c ie s
Location: _____________________________
N umbe r
Caug ht
N umbe r
M ar k e d
Nekton Sampling
Class Data Sheet
N umbe r
Re le ase d
Comme nts
FISH 221
Capstone I:
Team Building
Capstone I: Team Building
INTRODUCTION
As we begin our “capstone” field project, we will focus on developing teamwork, project planning,
self-esteem and self-confidence. We will also get a chance to practice our leadership, supervisory,
and following and listening skills. One excellent way to begin enhancing all of the above behaviors, attitudes and skills is to participate in the ROPES/Challenge course.
NOTE: ROPES is an incredibly effective process to develop teamwork.
PROCEDURE
Wearing a sturdy pair of shoes with good arch and ankle support is essential. Bring layered clothing
and raingear (NOTE: we won’t cancel if it rains or snows!).
You don’t have to be afraid of some of the more difficult challenges (i.e., the “pamper” pole and
“hole-in-space”). You can ask your team’s permission to excuse you from certain events. You’ll be
amazed just how much you will be able to accomplish, both individually and as a team!
LAB PRODUCTS
Following each event, teams will reflect on the significance of that event, either to us as individuals,
or as a team. These periods of reflection are definitely as important as the physical challenges
themselves.
There will be no write-ups due from this field exercise; however, you should reflect on this experience in the diary portion of your journal.
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NOTES FOR INSTRUCTORS
This activity is one of the first “capstone” activities we do, starting about midway through the
quarter. It provides opportunities for students to put into practive what they’ve learned thus far in
this course. This type of activity is recommended to help students pull together their collective
knowledge and apply what they’ve learned.
The ROPES Challenge Course is an example of an innovative team-building exercise. We’ve used it
with adult (instructor) groups as well as students, and responses are almost always positive.
These courses are available mainly through outdoor adventure centers throughout the country. Or
you may find this website useful, and a brief summary follows:
www.adventureropes.com/
Ropes Course
“We are a full service ropes/challenge course provider. We design, we build, we train, we teach. We
conduct inspections on existing challenge courses and we perform upgrades on those courses to
bring them into compliance with current safety standards. Our goal is to make the ropes course
experience available to everyone who is willing to accept the challenge. We believe that experiential
education is the most powerful tool ever developed to assist in learning.”
•FOR INSTRUCTORS•
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FISH 221
Capstone II: Ecosystem Study
Capstone II: Ecosystem Study
INTRODUCTION
This “capstone” will provide you the opportunity to put into practice all of the knowledge and
skills learned in previous classes, and specifically, those skills learned in the first five (5) weeks of
this course.
PROCEDURES
I. Project Planning for your Research Project
A. Brainstorming—Affinity and Tree Diagrams
B. Defining Scope-of-Work
C. Developing Tasks and Time Lines
D. Prioritizing Tasks
E. Assigning Personnel
1. Individual assignments
2. Team assignments
II. Project Management for your Research Project
A. Leadership and Supervision Skills
B. Teamwork Skills
C. Continuous Quality Improvement (CQI)
III. Project Completion and Delivery
A. Final Research Project
B. Oral Presentations
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Capstone
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NOTES FOR INSTRUCTORS
THE CAPSTONE RESEARCH PROJECT
I use a learning community approach in this project. I am including some notes on learning communities in this section. Included are guidelines for student project planning and development, and
presentation requirements.
A. SAMPLE RESEARCH REPORT OUTLINE
This is a sample of a research report structure that could be provided to students.
TITLE: Management Implications of Rockfish Tagging Research off the Southwest Washington
Coast
I.
Introduction
Begin with a general discussion of black rockfish; why tagging is necessary; past tagging efforts;
and end with the purpose (or focus) of this research report (NOTE: In a critical essay, this is your
thesis statement).
II.
Life history and habitat of black rockfish
A.
B.
III.
Importance of black rockfish
A.
B.
IV.
Life history
Habitat
To humans (commercial and recreational fishing)
To other animals (the food web)
Description of the study area
A.
B.
C.
Use a section of a coastal navigation chart
Photographs taken of coastline
Describe the coastal features and bottom characteristics of the area from USCG’s
Coast Pilot
V. Description of research vessel and gear
A.
B.
Interview a captain of a research vessel
Photos taken during interview
•FOR INSTRUCTORS•
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VI. Methods and materials
A. Personnel
1. WDF&W captain, crew and biologists
2. Volunteer fishermen
B. Equipment used to locate rockfish
1. LORAN and GPS
2. Radar and side scanning sonar
C. Gear and bait used to catch rockfish
1. Poles and reels
2. Bait and lures
D. The tagging process
1. Tags (photos and/or drawings
2. Tagging techniques (describe protocol; include data sheet as a figure)
VII.
Results and discussion
A. Compare past and present populations (use tables and graphs here)
B. Describe the current management plan for black rockfish in Washington State
C. Results of interviews regarding the effectiveness of this research with:
1.
2.
3.
4.
VIII.
The captain and crew
State fisheries biologists
A volunteer fishermen
Personal opinion (as an observer)
Summary and/or conclusions
A. Discuss how the tagging results will influence future management of black rockfish.
B. Summarize how important it is to include the public in a state research project.
IX. Bibliography or references cited
A. Journal reprints
B. Personal communications (interviews)
C. Articles from sport fishing magazines
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•FOR INSTRUCTORS•
X. Appendices
A.
B.
C.
D.
Tagging protocol used by state agencies
Actual data sheets taken from tagging trip
State management plans for black rockfish
Recreational fishing regulations for black rockfish
B. ORAL PRESENTATIONS OF RESEARCH REPORTS
I. Preparation
Your presentation should last no longer than 20 minutes. You should allow 3-5 minutes for
questions and answers. Your report should include:
a. an introduction (“tell them what you are going to tell them”)
b. the body of your talk (“tell them”)
c. a summary (“tell them what you just told them”).
NOTE: The format and organization of your presentation is just as important as its content!
AV equipment and supplies will be available for your presentation. Ask the instructor or assistant to
help you with these. They include:
a. white board (diagrams, drawings, etc.)
b. handouts
c. overheads (the instructor will provide)
d. slides
e. posters
f. Power-Point software
Useful tips:
¾ Remember to practice using AV equipment before your presentation.
¾ Practice your presentation in front of a mirror, your family, your roommate, your dog, or a
fellow student.
¾ Time your presentation so that you finish in 16-17 minutes, allowing 3-4 minutes for questions
from your audience.
¾ Note cards are acceptable, but practice enough so you only have to rely on them in a pinch. Do not
read from them!
¾ Always face your audience, even when talking from a slide, an overhead, or the white-board.
Relax and smile at your audience. Keep an eye on your audience to see if they are “in tune”
with your presentation.
¾ Don’t lean on the podium. Don’t confine yourself to the podium.
¾ Walk around, it will help you to relax.
•FOR INSTRUCTORS•
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STUDENT-DEVELOPED PROGRAM OBJECTIVES
1. To learn about and practice effective leadership, management and interpersonal relationship
skills—supervision, planning and organization, plan implementation, progress tracking, communications, coordination and cooperation.
2.
To gain confidence and critical skills through practice of all five elements of language—reading,
writing, speaking, thinking, and listening.
3.
To gain perspectives on local and global social, technological and natural resource issues from a
review of historical and contemporary writings, videos and discussions (seminars and guest
speakers). To be comfortable with answers to those questions regarding these issues that are
often incomplete and ambiguous.
4.
To learn about and practice state-of-the-art techniques related to:
¾
¾
¾
¾
¾
¾
¾
biological and chemical field and laboratory sampling and recording of data
experimental design and analysis of laboratory and field collected data utilizing
existing data base statistics and graphics software
literature searching
technical writing for local and state agencies using prescribed formats; using
advanced word processing of general and technically written letters, reports
grant writing
production of slide/tape, video and poster presentations
organizational work plans and progress evaluation
5. To learn the “art of seminaring”; especially, how to think and speak critically and ask good
questions of texts, teachers, our fellow students and ourselves.
6.
To gain perspectives on our own bias by considering the views of others.
7. To learn and practice the different stages of the writing process (general and technical)—
gathering materials, shaping it into writing, revising what one has written, and editing it for
publication and public presentation.
8. To practice working with others in exploring ideas, solving problems, and aiding others in
development of basic skills.
9. To experience delight, excitement, fear, rapture, astonishment, and a host of other emotions
that creative writing, thinking, and risk-taking can awaken.
10. To become better aware of and develop a personal framework of regarding ethics in the work
place, journalism, our scientific endeavors, and conservation of natural resources.
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•FOR INSTRUCTORS•
11. To explore, through works of literature and textbooks, fundamental questions of our
values, epistemologies, and lives in relation to technological change.
12. To learn to participate effectively and comfortably in an oral exchange of ideas about
significant literary works; to be able to listen and to formulate ideas spontaneously and
communicate them to others in large and small groups.
•FOR INSTRUCTORS•
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II
USING THE LEARNING COMMUNITY/LINKED COURSE APPROACH
The overall delivery of this research study will take on the nature of a “learning community,” either
as one which is done formally (team taught by two or more faculty), or informally, where two or
more faculty “link” their individual courses together (i.e., Aquatic Field & Lab Methods and
Technical Writing). Both will attempt to accomplish learning outcomes that jointly satisfy the
requirements of each of the individual courses. Students may be enrolled in just one or perhaps
both/all of the linked courses.
LEARNING COMMUNITIES—MARKETING AND ADVERTISING
Learning communities, if they involve team teaching of multiple courses, often require campus-wide
marketing and advertising. A poster or advertisement included in the college’s course catalog, or in
the local newspaper, is an effective way of getting the message out about the course. It is important
to inform and educate all divisions of the college (admissions, councilors, faculty, staff, administrators, etc.) about your learning community at least one quarter before it is taught. Planning by the
faculty co-instructing a learning community should start at least two quarters in advance. One-year
ahead of time is not too far ahead!
COURSE OBJECTIVES/TIME LINES
During the first week of the learning community, faculty and students will:
¾
¾
¾
Develop course objectives (requirements)
Develop a typical weekly schedule
Develop a corporate mission statement. The mission statement will continue to be
developed throughout the quarter.
Brainstorming is used extensively during the first two weeks to develop an affinity diagram, a tree
diagram and a set of time lines for the research study. These are all continuous quality improvement
(CQI) tools that allow student to practice management, leadership, teamwork, organizational,
facilitation, listening and following skills. Other quality management tools used are flow diagrams,
fish bone diagrams and multivoting (nominal group technique) for gaining consensus. The Memory
Jogger Plus + is an inexpensive pocket guide for students to learn these management and leadership
tools.
SEMINARS
Seminaring is a key ingredient of a learning community. One of the major goals of our learning
communities is to explore ecological and technological issues from various (i.e., organizational,
historical, economic, social and political) perspectives.
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•FOR INSTRUCTORS•
ASSESSMENT
Assessment is considerably different (and usually more comprehensive) in a capstone learning
community than a traditional class. Some of the assessment tools used include: a written selfevaluation by each student, an instructor-written evaluation of the student, comprehensive peer and
instructor evaluations, and a portfolio which includes all of the “end products” (learning outcomes)
including a final research report. This portfolio has proven very useful to the student/graduate for:
1) application to a 4-year college or university; and 2) applying for a job. It tends to provide a much
more comprehensive assessment of the student than just a grade on a transcript.
ORAL PRESENTATIONS
Students will make presentations to both general and professional audiences at the end of the field
exercise.
CHECKLIST FOR A SUCCESSFUL MEETING
Before the Event ~
_____
_____
_____
_____
Firm up the entire program well in advance; one to two months, if possible.
Be certain speakers know their topic and time requirements.
Outline program contents for publicity channels, including places and dates.
Complete planning for all physical facilities:
a. Meeting spaces; lunch and break areas.
b. Adequate exhibit spaces.
c. Registration supplies and personnel.
d. Audio-visual equipment and assistants.
e. Signs and bulletin boards for guidance of invited guests and speakers.
f. Invitations to guests, presenters, and the media.
_____ Line up community support and participation.
_____ Line up staff of hosts and hostesses.
_____ Print or copy brochures, programs.
_____ Arrange for photography and photographer.
During the Event ~
_____ Make sure someone is in charge; that everyone knows their responsibilities.
_____ Start sessions on time; announce at the beginning that this will be standard procedure.
_____ Keep sessions and speakers on schedule
•FOR INSTRUCTORS•
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Capstone
II
_____ Public address, recording and audio-visual equipment:
a. Have it ready and warmed up ahead of schedule; check that extra light bulbs are
available for overhead and slide projector; if using hi-tech. computer equipment, learn
how to use it, if passwords are required, etc.
b. Have qualified operators available.
c. Allow rehearsal time for all speakers to practice with A/V equipment.
d. Assign someone to turn lights off and on.
_____ Make a good photographic record of the program.
_____ Keep the program on schedule.
_____ Give credits to the planners and host/hostesses.
After the Event ~
_____ Personalized thank-you letters to speakers and assistants.
_____ Return all borrowed equipment and/or supplies.
_____ Pay all bills promptly.
SUMMARY
Since each capstone learning community is “student-driven,” there is no single outline to use as an
example. Students are allowed many choices during the quarter. Faculty must relinquish the
“control” or “authority” normally held in a traditional course. This does not mean, however, that the
students will not be productive. The opposite is most often true. Once the synergy of the
“community” kicks in (this often happens in the third to fifth week of the quarter, when students
finally accept the fact that they are 100% responsible for their own learning), near miraculous learning
occurs, and learning outcomes are accomplished within the prescribed time lines. All of this
requires much “risk taking” on the parts of both the faculty and students making up the community.
98
Capstone
II
•FOR INSTRUCTORS•
TEMPLATE FOR END-PRODUCTS
The following is a template with examples for constructing a list of “end product” requirements to
be completed by each student.
NOTES: Instructors can insert categories here of their own design, based upon the end-products
they decide upon. Students should be directed to pay close attention to specific due dates of each
end product.
End Products
Due date
Date Completed
Field notebook or journal
________
_____________
Written research report
________
_____________
Oral presentation of research
________
_____________
Demonstration of lab skills
________
_____________
Self evaluation
________
_____________
Instructor evaluation of student
________
_____________
Peer review
________
_____________
Gu i d e l i ne s for gra d i ng ora l
pre s e nt a t i ons of re s e a rc h
Cri t e ri a
P e rc e nt a ge s
O rga niza tion
20%
Content
20%
Use of AV ma teria ls
20%
Answers to Questions
20%
Public Spea k ing Sk ills
20%
Tota l
100%
•FOR INSTRUCTORS•
99
Capstone
II
100
Capstone
II
Se m inar
Lab/
Pr oje c t
Fie ld
PlanPar tic i- ning
pation
Te am wor k
Le ade r - Re se ar c h
ship
Re por t
(1st
D r aft)
Re se ar c h
Re por t
(Final
D r aft)
N ote s
To students:
Please rate yourself and your teammates performance with regard to the following area zero (0) being the lowest, five (5) the highest.
Your ratings will be kept strictly confidential.
N am e
SELF-ASSESSMENT PEER REVIEW OF PERFORMANCE
101
Capstone
II
N am e
Se m inar
Lab/
Pr oje c t
Fie ld
PlanPar tic i- ning
pation
Te am wor k
Le ade r - Re se ar c h
ship
Re por t
(1st
D r aft)
INSTRUCTOR EVALUATION OF STUDENT’S PERFORMANCE
Re se ar c h
Re por t
(Final
D r aft)
N ote s
REFERENCES
Brassard, M. 1989. Memory Jogger Plus +. First Edition. GOAL/QPC. 13 Branch Street, Methuen,
MA 01844.
Department of the Navy. 1992. Handbook for Basic Process Improvement. Chief of Naval Operations
Executive Steering Committee and the Department of the Navy’s TQL Office, Washington, D.C.
102
Capstone
II
FISH 221
WEEKLY SEMINAR ASSIGNMENT #1
Weekly Seminar Assignment #1
Definitions of Ecosystem Management and Related Terms
Available on the Web at: classes.aces.uiuc.edu/NRES325/defin.html (Pages 1-8)
Answer the following questions regarding our seminar reading for this week. Write your answers on
separate sheets of paper. You may also find it useful to jot down comments, questions, and page
numbers on a separate sheet to bring to seminar.
1. What did Aldo Leopold and Eugene Odum say about ecosystem management? (pg 1) Roughly,
what year(s) did each of these “visionaries” make these statements?
2. Write down, sentence by sentence, the entire definition of ecosystem management authored by
the Federal Interageney Ecosystem Management Task Force (June 1995). After each sentence, write
in your own words, what you think the Task Force meant by each statement. Continue until you
have completed the definition. (pg 1)
3. What else does the Task Force’s approach emphasize? (pg 1)
4. List and discuss the seven (7) elements that make up ecosystem management (pg 2) according to
the Ecological Society of America’s (ESA’s) “The Scientific Basis for Ecosystem Management
(1995).”
5. Summarize in one paragraph, what the ESA says about ecological science as a basis for ecosystem
management. (pp 2-3)
6. What are some basic differences between the positions taken on ecosystem management between
the U.S. Fish and Wildlife Service and the U.S. Forest Service? (pg 4)
7. Differentiate between ecosystem “health” and ecosystem “integrity.” (pg 5)
8. List several ecosystem heath goal, objectives and indicators. (pg 6)
9. What are the five (5) main ecosystem management goals listed by Edward Grumbine?
10. What is biotic integrity according to James R. Karr?
103
Weekly
Seminar 1
FISH 221
WEEKLY SEMINAR ASSIGNMENT #2
Weekly Seminar Assignment #2
“Learning to Work Together”
Reference Material: The Team Handbook—How To Use Teams To Improve Quality. 1992. 18th Printing.
Joiner Associates, Inc. Madison, WI.
You will be responsible for the contents of Chapter 6 of the “Team Handbook.” No formal written
questions will be required; however during seminar time you are required to reflect upon and write
your responses to Question #1 (below) in the Diary Section of your journal.
Seminar (First Session)
1. After reading and studying Chapter 6 of the Team Manual thoroughly:
a.
Where do you think your team is at this time regarding: forming, storming, norming or
performing?
b.
If your team has progressed past the forming stage, describe the behaviors that existed at
earlier stages, and those behaviors that exist now (15 minutes for this exercise). Record
behaviors at each stage on flip chart.
c.
What behaviors describe your team at your current stage of team development?
2. Discuss the ten (10) ingredients for a successful team (pp 6-11 to 6-21) regarding:
104
Weekly
Seminar
2
a.
Indicators of potential trouble. Which of these trouble indicators apply to your team?
Brainstorm around your circle of members, having each team member offer one trouble
indicator (that applies to your team) per round (no criticizing or discussion for this
exercise; it is okay to pass). Facilitator should keep this moving fairly quickly. Recorder
should write responses on a flip chart. Be sure however, that everyone gets a chance to
get all of their “indicators” out in the open, and up on the chart. Go around the circle
as many times as needed to complete this (about 20 minutes for this exercise).
b.
Brainstorm again, addressing the question “What can I do to help my team progress to
the performing stage?” If you are already at the performing stage, address the question
“What can I do to make our team perform even better?” Each member should begin
his/her statement with: “I can help our team be more successful by . . .” Again, the facilitator
should keep this moving, and the recorder should use a flip chart (allow about 20
minutes for this exercise).
Seminar Continues (Second session)
3. Teams get back together. Reporters use flip charts to report to the entire class. (Allow 10
minutes maximum for each team report. Time keepers—remind facilitator two minutes before
time is up).
4. Class evaluation of this seminar exercise (10 minutes).
105
Weekly
Seminar 2
FISH 221
Quiz I
Quiz I
1. Define the following terms and list a living representative of each (be as specific as possible).
a. plankton
b. periphyton
c. macrophyton
d. macroinvertebrates
e. autotrophic
f. heterotrophic
2. Explain the significance of each of the following as indicators of polluted and/or clean water.
a. plankton
b. periphyton
c. macrophyton
d. macroinvertebrate
3. List the five (5) essential pieces of information to be included on sample labels that are inserted
into sample containers immediately as plankton samples are collected.
106
Quiz 1
FISH 221
Quiz II
Quiz II
1. Define the following biometrics terms:
a. statistic
b. biometrics
c. parameter
d. standard deviation
e. observation
f. mode
g. character of interest
h. universe
i. experimental unit
j. population
k. sample
l. frequency histogram
2. What is a normal distribution curve? Why are comparisons between our frequency polygon and
the normal distributions curve useful?
3. List two (2) types of statistics that describe central tendency and two (2) that describe measure
of dispersion. Briefly explain what they are.
107
Quiz 2
4. Consider the following sample set of numbers: 2, 3, 3, 4, 4, 4, 8, 10, 12
a. Construct of frequency table using the above data.
b. Find the range.
c. Calculate the mean.
d. What is the median?
e. What is the mode?
f. Average deviation?
g. Variance?
h. Standard deviation?
i. Draw a frequency polygon using the data (sample set of numbers) provided.
108
Quiz 2
FISH 221
Sample Mid-Term Exam
Sample
Mid-Term Exam
(100 points)
Answer any five (5) of the following. (use a spearate sheet of paper).
1. Explain how you would use the backpack electroshocker to sample fish populations in a creek.
Be specific about start-up and shut-down procedures, voltage and wattage settings, type of
current used and why, and what water quality measurements must be made prior to use of
electroshocker? Why? What precautions should be taken to avoid undue stress to fish?
2. Describe in detail the use of seines, trawls, chemicals, and hook and line fishing as active fishing
methods. Give advantages and disadvantages of each method and tell when each might be the
preferable method over the others.
3. Describe two (2) types of entanglement devices and two (2) entrapment devices used in passive
fish sampling. Be specific about how, when, and where used. Compare advantages and
disadvantages.
4. Define plankton, periphyton, macrophyton, and macroinvertebrates. Be specific about the types
of plants or animals that make up these major groups and where they might be found in the
aquatic or marine environment. How do they serve as an “indicator” of pollution? Again, cite
specific examples.
5. Define five (5) of the following biometric terms:
a. experiment
b. observation
c. population
d. sample
e. character of interest
f. universe
g. experimental unit
h. parameter
i. statistic
6. Explain how you would determine the “standing plankton crop” or “biomass” (both qualitatively
and quantitatively) in a lake. Be specific about equipment and techniques used in sample
collection and analysis.
109
Sample
Midterm
7. Your assignment is to research the change in the periphyton communities related to the
installation of a secondary treatment plant on the _________ River. List all of the major and
incidental observations that should be included in your field notes. You collect several
periphyton samples to analyze in the laboratory. What information should be included in your
field notes? What information should be included on the sample label you insert in the
collection container?
8. List at least five (5) major types of observations to be included in field notes when collecting
plankton or fish in a lake. Also list at least five (5) pieces of information to be included on labels
to be inserted into sample container.
9. What is ecosystem management? What is adaptive management? In your discussion compare
the way we have managed natural resources in the past with these new management philosophies, strategies and practices.
110
Sample
Midterm
FISH 221
Oral Exam and Study Guide
Sample
Oral Exam and Study Guide
1. You will be asked to verbally define and discuss five (5) of the terms from the attached list of
watershed related terms. The instructor will randomly select terms, so you’ll have to know them all.
2. Describe (in detail) each of the following physical, chemical and biological sampling procedures.
Describe in terms of equipment needed, protocol for sampling and analysis, and the nature of the
data collected (units of measurement, description of data sheets, use of data):
•
•
•
•
•
Plankton sampling and analysis
Surber sampling
Otter trawling
Beach seining
Flow (water velocity or discharge)
3. Be prepared to discuss benefits (or disadvantages) of Continuous Quality Improvement (CQI) as
a set of project planning and management tools to carry out our research study this quarter (i.e., did
they help or hinder our goal to complete several major team “end products?”). You will be graded
strictly on how well you defend your point of view (pro or con); not on what you think I want to
hear.
4. Be prepared to discuss pros or cons of the “learning community” (teamwork, seminars, students
teach other students, journals, oral presentations, portfolios) approach to learning versus more
traditional approaches. Again, you will be graded on how well you defend your point of view; not
on what you think the instructor wants to hear.
111
Sample
Oral Exam
Study Guide
Watershed Related Terms
The following is a list of watershed-related terms that will be “fair game” for definitions and
description for the final oral exam:
Accuracy
Adaptive management
Alkalinity
Baseline survey
Bed load
Benthic
Biodiversity
Bioregion
Buffer strips
Canopy
Channel forming discharge
Community
Conductivity
Coniferous
Control
CQI (Continuous Quality Improvement)
Cover
Cubic feet per second
Cumulative effect
Debris flow
Deciduous
Discharge
Dissolved load
Dissolved oxygen
DOM (Dissolved Organic Matter)
Ecosystem
Ecosystem management
Ecotone
Electroshocking
Ephemeral
Erosion
Experimental design
First order stream
Flood plain
Fluvial
112
Sample
Oral Exam
Study Guide
•FOR INSTRUCTORS•
Geomorphology
GIS (Geographic Information Systems)
Glacial flour
GPS (Global Positioning Systems)
Gradient
Gully erosion, gullying
Habitat units
Hardness
Hydraulic permit
Intermittent stream
LWD (Large woody debris)
Limiting factor
Macroinvertebrates
Mass wasting
Meander
Non-point source pollution
Parameter
pH (hydrogen ion concentration)
Plunge pool
Point source pollution
Pool
Population
Precision
Reach
Replicate
Riffle
Riparian zone
Scoured
Side Channel
Stream segment
Stream typing
Surface erosion
Suspended sediments
Synergistic
Tailout
Thalweg
Time lines
Total solids
TQM (Total Quality Management)
Watershed
Wetlands
NOTE: Portions of the study guide can be selected by the instructor and used as the oral exam.
113
Sample
Oral Exam
Study Guide
FISH 221
Sample Final Exam
Sample
Final Exam
Definitions
1. Define and give “real world” examples of the following biometrics terms.
a. observation
b. experimental unit
c. universe
d. character of interest
e. biometrics
Fill-Ins.
1. Parameters describe ________________ while statistics describe ________________. An
experimental unit is to the ________________, as a sample is to the ________________.
2. A term that indicates how many times a particular score occurs in a collection of data is known
as the ________________.
3. The difference between the highest and lowest values in a distribution of values is called the
________________, while the square root of the sum of squares of the deviation of the values
from their mean, divided by the total number of observations minus one (1), is called the
________________.
114
Sample
Final Exam
4. The arithmetic average of all values in the sample distribution is the ________________; the
value that occurs most frequently is the ________________; and the midpoint of the distribution
is the ________________.
Short Answers.
1. List two (2) types of statistics that describe central tendency and two (2) that describe measure of
dispersion. (Briefly tell what they are)
2. What is a normal distribution curve? Why are comparisons between our frequency polygon and
the normal distribution curve useful?
3. How is standard deviation useful to the biometrician?
In your own words, list five (5) of the ten (10) principles for conducting biological sampling. Cite a
“real world” example of each.
115
Sample
Final Exam
Definitions. Define the following terms and list a living representative of each (be as specific as
possible).
a. plankton
b. periphyton
c. macrophyton
d. macroinvertebrates
e. autotrophic
f. heterotrophic
Short Answers. Explain how each of the following are of significance as indicators of polluted and/
or clean water.
a. plankton
b. periphyton
c. macrophyton
d. macroinvertebrate
Essay. Answer any five (5) of the following.
1. List five (5) different physical, chemical and/or biological factors that should be controlled in a
bioassay and explain why each is important to the final outcome of the experiment.
2. Describe in detail two (2) types of active fish sampling methods or devices, and two (2) types of
passive methods or devices. Be specific about how, when, where, and why used. Give the
advantages and disadvantages of each.
116
Sample
FInal Exam
3. Describe how you would design a bioassay to determine the toxic effect of cupric ion (Cu++)
on coho salmon in the Chehalis River. When might a static bioassay be appropriate? A flowthrough system?
4. Explain the principal of electroshocking and how you would use the backpack electroshocker to
sample fish populations in a creek. Be specific about start-up and shut-down procedures,
voltage and wattage settings, type of current used and why, and what water quality measurements must be made prior to use of the electroshocker.
5. Explain how you would determine the “standing plankton crop” and “biomass” (both qualitative
and quantitative) in a lake. Be specific about equipment and techniques used in sample
collection and analysis.
6. List at least five (5) major types of observations to be included in field notes when collecting
plankton or fish in a lake. Also list at least five (5) pieces of information to be included on label
to be inserted into sample container.
7. Using the ten (10) principals of experimental design, describe how you would formulate your null
hypothesis (describe each step in getting there) and how you would test whether bait type made
any difference in catches of spot shrimp in Puget Sound.
117
Sample
Final Exam
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