1 Animal Behaviour: Biology 3401
Laboratory 2: The effects of physical and biological factors on various
invertebrate organisms
Parts of this lab were taken from or adapted from: Glase, J. C., M. C. Zimmerman, and J. A. Waldvogel. 1992.
Investigations in orientation behavior. Pages 1-26, in Tested studies for laboaratory teaching, Volume 6 (C. A.
Goldman, S.E. Andrews, P.L. Hauta and R. Ketcham, Editors). Proceedings of the 6th Workshop/Conference of the
Association for Biology Laboratory Education (ABLE), 161 pages.
Questions to Prepare You for this Laboratory
• What are three types of orientation stimuli that can be sensed by some organisms but
not humans?
• What characterizes a taxis orientation response?
• What characterizes a kinesis orientation response?
• Is the manner in which you would locate the source of an odor in a dark room a taxis, a
kinesis, both, or neither?
• Is the manner in which you would locate the source of a sound in a dark room a taxis, a
kinesis, both, or neither?
Introduction
Humans have been aware of animal orientation, migration, and navigation for
thousands of years. Whole civilizations have thrived or perished based on their
understanding of the movements made by principal animal food sources. Yet, it was not
until early in the 20th century that a rigorous analysis of orientation mechanisms began,
when our knowledge of sensory systems and how other animals detect the world
improved considerably.
Orientation refers to the spatial organization of movements. Since movements are
elements of behavior, orientation and behavior are intimately associated. For simplicity,
we will define behavior as any overt manifestation of life by an animal, especially one
that takes the form of movements. A behavior pattern is the fundamental unit of
behavior, and is defined as a sequence of movements characterized by a specific
configuration in time and space. This underscores the special significance that spatial
organization has for behavior. Every behavior is spatially oriented in some way. Whether
an animal walks, grooms, catches prey, or interacts with a social partner, "where" and "in
which direction" are indispensable features of its behavior pattern. Thus, we can define
orientation as the process that animals use to organize their behavior with respect to
spatial features.
The specific orientation systems used by an animal correspond to the features of
its environment. Many terrestrial organisms are sensitive to humidity levels, and are
therefore capable of orienting with respect to moisture gradients. But humidity is an
environmental feature that is not relevant in a totally aquatic habitat, and as a result
animals that live in water must use physical gradients based on other parameters (e.g.,
temperature or salinity) to help direct their movements. Some orientation stimuli are
available to both terrestrial and aquatic organisms; these include gravity, light and
2 magnetism.
This laboratory exercise will allow you to investigate orientation behavior in a
variety of animals. It will help you understand the scientific method by giving you
experience in conducting and interpreting data involving animal orientation. Orienting Stimuli In orientation studies, one first attempts to identify the nature of the stimuli to
which the animal is orienting. Light, gravity, sound, and mechanical stimuli, as well as
temperature, chemical, and moisture gradients are all likely candidates. As with a moth
flying into a candle, the nature of the orienting stimulus may be clearly apparent.
However, if the animal is orienting to a stimulus for which humans have no receptor
organs, identification of that stimulus will be much more difficult. Orientation to
ultraviolet and polarized light, magnetism, electrical fields, and some acoustic stimuli are
of this sort. Frequently, organisms respond simultaneously to several stimuli while
orienting. Thus, one must be cautious in interpreting observations of orientation behavior
since the stimulus most obvious to human senses may not be the most important factor
determining the animal's behavior. Frequently an orienting stimulus also elicits a
behavioral response. For example, in prey-catching and courtship behavior, the animal
often first orients toward the prey or mate, then performs the appropriate behavior to
capture the prey or attract the mate. The presence of the prey or potential mate in the
environment causes the animal to orient appropriately as well as to perform other
behaviors. At the same time that an animal is stalking prey or courting, it is also using
gravity as a stimulus for body orientation relative to the earth. As a tuna pursues a
mackerel in the open ocean, the mackerel elicits and orients the predatory behavior of the
tuna, but gravity and light stimuli are also used by the tuna for general body orientation.
In addition to species differences for a given orientation behavior, the nature of
the orienting stimulus itself may vary as a function of the animal's age. Many nestling
birds, for example, show a gaping response which elicits parental feeding. When the
nestlings first hatch they are blind, and the gaping response is released by mechanical or
auditory stimuli provided by the parent birds. The nestlings gape vertically, with gravity
being the main orienting stimulus. Later, after a nestling can see, the sight of the parent
bird not only elicits the gaping response, but also orients it. Classification of Orientation Responses The ways that animals orient to their environment are diverse, and certain
schemes have been developed to classify these responses in reference to underlying
similarities. The classification system presented in this laboratory was first suggested by
Fraenkel and Gunn (1961 - The Orientation of Animals). Kinesis One important distinction that Fraenkel and Gunn make depends on whether the
animal's body is oriented with respect to the stimulus source. A movement that does not
involve orientation with reference to a stimulus source is known as a kinesis where the
stimulus produces either a change in the speed of the animal's movement (orthokinesis)
or in the animal's turning rate (klinokinesis). These two responses effectively change the
3 position of the animal with respect to the stimulus source. Several examples should
clarify this point.
Isopods (terrestrial crustaceans) prefer moist habitats. In some species, as the
relative humidity of the environment increases, the amount of time the animal is
stationary also increases. This response tends to keep an isopod in damper areas. As
another example, some insects cannot detect the direction of an odor gradient, but their
rate of locomotion varies with the strength of the odor. Thus, if an insect moves rapidly at
low concentrations of a chemical and slowly at high concentrations, it should eventually
arrive at the source of the odor. The human body louse (Pediculus corporis) finds its host
by a kinetic response to a number of stimuli including temperature, humidity, and odor.
When in a favorable environment with respect to these stimuli, the louse travels in
straight lines. However, if it encounters an unfavorable environment, it turns until a
favorable environmental zone is once again encountered. In summary, a kinesis involves
quantitative variations in an animal's speed or turning rate with no fixed orientation of the
body relative to the stimulus source. Taxis
In a taxis, the animal's body is oriented in some linear manner relative to a
stimulus; either directly toward it, directly away from it, or at a fixed angle to it.
Locomotion may or may not be involved in a taxis. This kind of response may be shown
for light, heat, moisture, gravity, sound, chemicals, or other stimuli.
For the next two weeks you will be examining the effect of various physical and
biological stimuli on the behavior of a variety of invertebrates.
Materials and Methods
A. Physical Factors
1. Humidity and Light Preference in pillbugs
These terrestrial crustaceans, sometimes called pillbugs (Armadillium sp.), are
common inhabitants of leaf litter and soil. They feed on decaying organic material as well
as algae, moss, and bark. Isopods have a pair of compound eyes, two pairs of antennae
(although only the second pair is prominent), and seven pairs of legs. When disturbed or
desiccated they will roll up into a ball, looking rather pill-like.
Experiment one: This experiment will be conducted in an elongated Plexiglas chamber
that can be used to collect quantitative data on the response of individuals to light,
humidity, or a combination of these stimuli (Fig. 1). At one end there is anhydrous CaCl2
(a desiccant) and a wet paper towel in the other end. This sets up a humidity gradient that
we are assuming is changing consistently over the length of the apparatus. For this part of
the experiment, observe the animals under red light (most invertebrates have a lower
sensitivity to red light). Place 10 individuals in the chamber via the central stoppered
hole (be careful not to introduce debris into the chamber). Every minute count the
number of pillbugs in each section of the chamber (there are paper sheets to place under
the chamber to indicate which section the bug it in).
4 Enter the data on the sheet on the next page (Data table 1). Continue counting for 30
minutes.
Experimental Setup for Pillbug Choice Experiments
Introduce pillbugs here
Desiccant
(CaCl2)
Moistened
paper towel
1
Dry
2
3
4
5
6
7
SECTION
8
9
10
Humid
5 Data table 1 – Data Table for Pillbug Wet vs. Dry Choice Experiment
Section
Observation
#
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
26
27
28
29
30
Mean
S.D.
Dry
2
1
3
4
5
6
7
8
9
Humid
10
6 Experiment 2. This experiment is essentially a repeat of Experiment 1 but testing the
response of pillbugs to light vs dark. Place the black plexiglass covers over one half of
the chamber to create the dark end. Introduce 10 pillbugs into the chamber as before and
wait one minute before beginning observations. After each minute, count the number of
animals in the light sections (to get the number in the dark you have to count the number
in the light section and subtract from 10…). Enter the data in Data Table 2. Continue
counting for a minimum of 10 minutes and stop when the number of animals in one
section is 0 for three successive minutes.
The Interaction of Factors
After you have gained some insight into the pillbug's response to humidity and
illumination independently (and have collected enough data to support your views!), you
can examine the interplay between these two stimulus types. Is the response to humidity
changed under conditions of high illumination? What if the individuals are offered a
brightly illuminated, humid environment versus a dark, dry environment?
Experiment 3
In this experiment, you will be looking at the interaction between humidity and
light in the response of pillbugs. Set up the experiment as in experiment 1 but put one
end of the chamber (either the humid or dry end) into the opaque chamber. Introduce ten
pillbugs into the chamber and wait one minute before beginning observations. After each
minute, count the number of animals in the dark and in the light sections. As before,
continue counting for a minimum of 10 minutes and stop when the number of animals in
one section is 0 for three successive minutes. Enter the data into Data table 3.
Repeat the experiment but this time switch the end the chamber that is in the
opaque chamber. Introduce ten fresh (i.e. untested) pillbugs into the chamber and repeat
the experiment as above.
Experimental Setup for Pillbug Combined Choice Experiments
Desiccant or
Moistened paper
towel
Introduce pillbugs here
Opaque
chamber
1
2
3
4
5
6
7
8
SECTION
9
10
Desiccant or
Moistened paper
towel
7 Data table 2 – Data Table for Pillbug Light vs. Dark Choice Experiment
Section
Observation
#
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
26
27
28
29
30
Mean
S.D.
Dark
Light
8 Data table 3 – Data Table for Pillbug Combination Factors Experiment
Observation #
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
26
27
28
29
30
Mean
S.D.
Dry -Dark
Humid – Light
Dry – Light
Humid - Dark
9 2. Geotaxis in snails/slugs
Background:
Gastropods (including limpets, whelks, slugs and snails) generally move slowly
and ‘sluggishly’. For that reason, their locomotion is relatively easy to study. Under
certain conditions they move consistently in one direction, and are apparently welloriented with respect to gravity. Also, the direction of their movements often seems to be
influenced by the paths taken by other nearby snails.
This portion of the laboratory concerns the study of three aspects of the
locomotory behaviour of a terrestrial snail Helix aspersa: (1) undisturbed movements, (2)
gravity responses, and (3) the tendency to follow trails left by other individuals.
Helix aspersa is commonly found in gardens, hedges and woodlots. This snail is
normally inactive during daylight and under dry conditions, and moves and feeds
especially at night or after rain. It feeds on living vegetable material, from which
fragments are abraded by means of a toothed radula. In winter and during prolonged dry
periods, Helix seals up the aperture of its shell with a film of mucus hardened with
calcium phosphate.
Many, if not all snails, will be quiescent at the start of the laboratory session and it is
essential to stimulate them and have them moving about actively immediately before use.
This is best done by immersing the snails in tepid (lukewarm) water for a few minutes.
As soon as a snail shows signs of activity it should be removed from the water, dabbed
dry and introduced to the experimental set-up.
Handle the snails gently, especially when lifting them up from an attached position.
They are not likely to move in a natural, consistent way if you roughly tear them loose
from a substrate.
PART 1: Movements in relation to gravity (geokineses/geotaxis) are to be studied as
follows. If Helix aspersa (the land snail) is available use Helix. Otherwise, use the slugs
available in lab.
• Place a fresh slug on the centre of a clean plexiglass plate that has a grid of squares
drawn on the reverse side. Clamp the plate in a vertical position with one edge on
the bench top (angle of inclination=90o). As soon as the slug begins moving, start
a stopwatch, and plot its course to scale on a separate sheet of graph paper. After
5 minutes, or when the slug has reached the edge of the glass plate (whichever
occurs first), remove it. Determine the angle of orientation of its path with respect
to the side of the plate in contact with the bench. Do this by drawing a straight
line on the graph paper that represents as accurately as possible the direction of
the slug’s path. If a sharp change in direction occurred during the test, two
separate angles of orientation should be determined.
• By using a piece of string to estimate the distance travelled by the snail, and the time
between start and end of the test, determine its speed of movement (cm/min).
10 • Repeat the above procedure with two more slugs (thus yielding 3 replicates), being
sure to wash the glass plate thoroughly after running each animal.
• Then change the angle of inclination of the plane to 45o from the horizontal and repeat
above procedure with the same three slugs (or fresh slugs if the supply is large
enough).
• Finally, repeat the steps again with the plate at 75 and 25o inclination. 3 slugs each for
a total of 12 separate tests.
•
• Record all your data in Data Table 4.
• Prepare a graph showing the mean angle of orientation and another graph showing the
mean speed of movement of your slugs in relation to the angle of inclination of
the plate. Each mean value should be based on 3 replicate measurements.
• In discussing your results, consider the following: What are the sensory structures that
enable a slug to detect changes in gravity? Where are they located? What are
some environmental variables that could affect the behavior of slugs in this
laboratory exercise? What might be the significance of the observed responses to
the slugs in nature? Are they taxes or kineses?
Experimental set-up for snail geotaxis experiment
11 Data Table 4: Data table for slug geotaxis experiment
Plate angle
Slug #
Slug path angle
Speed (cm/min)
(degrees)
(degrees)
90
1
90
2
90
3
45
45
45
1
2
3
75
75
75
1
2
3
25
25
25
1
2
3
Distance (cm)
PART 2: Chemoreception in slugs (trail following)
• Purpose: to study trail following by determining the degree to which the mucous trails
of a ‘blazer’ and a ‘follower’ snail coincide.
• The tendency for one animal to follow the recently laid mucous trail of another
conspecific has been demonstrated in several gastropods, including terrestrial
snails. Examine trail following in slugs as follows.
• Method: Place one active slug (the ‘trail blazer’) at the centre of a piece of clean
plexiglass lying flat on the bench top. Plot to scale the path of this slug until it has
moved 15-20cm (within 5-8 minutes). Then remove it and place a second slug
(the ‘follower’) at the same starting point, and oriented in the same direction as
the blazer. Observe and plot the path of the follower on the same graph paper until
it has moved at least as far as the blazer did. Remove the follower and determine
the coincidence index (C.I.) of the two mucous trails using the following
equation:
C.I. = Lc/(Square root of (Lb X Lf))
Where Lc is the length of the follower trail that coincides with the blazer trail, Lb is
the length of the blazer trail and Lf is the length of the follower trail. Use string to
estimate each of these distances.
12 • Repeat the above procedure with two more pairs of slugs, making sure that the glass
plate is washed between tests. Summarize your data in a table (Data table 5),
showing values of Lc, Lb, Lf and C.I. for each pair of slugs and an overall mean
(SD) C.I. Discuss the possible functional significance of trail following in a slug.
Data table 5: Data from trail following in slugs
Trial #
Lc
Lb
1
2
3
Mean
SD
Lf
CI
3. Chemoreception in flour beetles (Tenebrio molitor; effect of pheromones on mate
attraction)
•
See additional hand-out.
•
ANIMAL BEHAVIOUR: BIOLOGY 3401
Part of Lab 2: Physical and Biological factors (additional hand-out)PHEROMONES AND MATE ATTRACTION IN THE FLOUR BEETLE,
Tenebrio molitor*
INTRODUCTION:
Many insects possess extremely well developed olfactory organs and in
consequence, rely upon chemoreception to direct such biologically essential activities as
location of food, mate or oviposition site, induction of copulatory behaviour and alarms
signaling the approach of a predator. As well, exchange of chemical information between
insects of the same species can, in some cases, bring about physiological changes that
result in modification of the insect’s pattern of development.
The term pheromone was used by scientists in the late 1950’s who were studying
chemical communication in insects. They applied the term to any compound that is
released by one member of a species to carry information to another of the same species.
Studies on gypsy moths showed just how strong the physiological response to these
13 chemicals can be. They were able to fool male moths into copulating with anything from
oak leaves to filter paper by simply spraying the surface with the female pheromone.
Extensive work has been done on insect pheromones, including studies on the
specificity and degree of response to natural and synthetic pheromones. Studies show
that for the purpose of mate location, usually the female (less often, the male) releases a
pheromone to attract the opposite sex. In some insects, copulatory behaviour following
mate location is induced by an aphrodisiac pheromone released by either the male or
female, depending on species.
Copulation though is not the sole purpose for pheromone release in any species.
Other examples of pheromone use include: attraction to a feeding site (e.g. trail laying in
ants); alarm pheromones, produced when a social insect is threatened in some way;
control of long-term developmental processes (e.g. maintenance of caste structure in
termite colonies). In most insects, pheromones are produced by epidermal cells
compacted into discrete glands and concentrated in defined areas beneath the cuticle.
* Adapted from: Physiology Laboratory Manual, 1993, T. Rand, Saint Mary’s
University.
Most pheromone communication is accomplished by mixtures of compounds
(‘multicomponent pheromones’ or ‘pheromone blends’) and it is the relative proportion
of constituents comprising the blend (as well as the sensitivity of the receptors of the
receiving individual) that differs among species and confers an element of specificity to
the response.
An industry is being developed around the potential use of pheromones for insect
control purposes. Pheromone-baited traps can be used to: (a) monitor the presence of a
pest in the ecosystem, to determine whether or not it would be an appropriate time to
apply insecticides; (b) remove large numbers of pests directly from the ecosystem (masstrapping); (c) confuse natural pest populations by saturating the atmosphere with
pheromone and, thus, preventing mate location.
In today’s laboratory, you will be studying sexual attraction via pheromones in
one insect: the meal worm beetle (also known as the flour beetle), Tenebrio molitor.
14 You will examine the capacity of female insects, and extracts prepared from virgin
females, to attract males and also the female response to live males and to male extracts.
METHODS:
A. General:
Because Tenebrio is photosensitive and actively avoids the light, the entire
laboratory will be carried out under red light to which the beetles are less sensitive. As
you obtain the data for your group, enter it in the Tables provided and on the blackboard.
Today’s laboratory relies heavily on class data. Tenebrio were sexed and segregated as
pupae. Pheromone extracts of Tenebrio were prepared recently.
There are two parts to this lab – assessing the male response to female
pheromones and the female response to male pheromones.
A. Male response to female pheromones
For each trial, you will be using a Petri dish, the bottom of which is lined with a
sheet of filter paper. This filter paper must be changed for each trial. Put a single male
in the Petri dish and allow him 5 minutes to acclimate. Take a glass rod from the ethanol
and dry it off by repeatedly shaking it and wiping it with a Kimwipe tissue. Avoid
handling the tip of the rod with your fingers. Dip the end of the rod in the experimental
fluid and let it air dry for 30 seconds. Place the rod centrally on the filter paper in the
petri dish and replace the cover. Record the response. If there is no response in 5
minutes, terminate the trial and get a new male.
To rate the male response, use the following scale:
0 - no attraction
1 - some attraction, circles the glass rod
2 - some attraction, touches the glass rod, more than once
3 - more attraction, stays on the glass rod, or repeatedly returns and
touches it; moves along the length, keeping contact with it
4 - attempts copulation or shows copulatory behaviour
15 Response in the male upon perception of a female begins with the beetle waving
his antennae. He will then extend his prothoracic legs, resulting in raising his body, the
rearing and antenna waving becoming jerky and faster. Copulatory response involves the
male, once in contact with the female, stroking her with his antennae, then climbing up
on her quickly while continuing to stroke her sides with his front legs. He will then bend
his abdomen down and around the end of the female (or the rod) to make genital contact.
He continues to beat against her back with his antennae during copulation.
A positive
behavioral response may also be displayed by the beetle crawling along or staying affixed
to the rod, making no attempt to leave it.
You will be testing the males with several different solutions. These are:
1. an untreated glass rod (control)
2. female pheromone extract (A, B, or C) (these solutions are of graded
concentration expressed as number of insects extracted/ml, solutions are: A, 2.5/ml; B,
0.625/ml; C, 0.156/ml)..
3. bornyl acetate (a female cockroach pheromone)
In each test, you should note which level of response that the male attained on the scale
above.
B. Female response to the male pheromone
Female response is far more passive than the male response and thus must be
rated differently. When a female detects a male pheromone, she tends to slow down but
she does exhibit some rearing and antenna waving. Studies have shown that females will
gather in areas where male pheromone has been placed. We will use these two criteria to
gauge female response.
We are using chambered models (Fig. 1) to determine response in the female.
Two virgin females are placed in each of the six outer chambers and allowed 30 minutes
to acclimate. At 15 and 30 minutes in the acclimation period, examine the activity and
distribution of females in the chamber. To score the distribution, divide the entire
chamber into two sections (near and far zones, as defined by concentric circles (see Fig.
1). Note the total number of females in the near and far zone in each chamber.
16 To score the activity, record the number of seconds each female spends walking
in a 30 sec. observation period.
After 30 minutes, place one of the following stimuli in the central chamber of the
apparatus.
•
a live male – WEDNESDAY LAB
•
a rod dipped in male extract – WEDNESDAY LAB
•
a clean glass rod (control) – THURSDAY LAB
•
a rod dipped in female extract - THURSDAY LAB.
You will now be monitoring the chamber for 30 minutes. Every 3 minutes note the
position of the females (i.e. whether each is in the near or far zone). At 10, 20, and 30
minutes record the activity as above.
far zone
near zone
central chamber
Fig. 1. Top view of the choice chamber used in female choice experiment.
Each section of the chamber is divided into a near and far zone to assess the
attraction of females to the stimulus in the central chamber.
STATISTICAL ANALYSES
The part of analyzing this lab that will be the most challenging is the statistics to use
when assessing the response of males. Male response has been scored qualitatively (as
opposed to quantitatively) meaning that the responses have been rated on a scale from 0
to 4. These kinds of data are called ordinal and must be analyzed using non-parametric
17 tests. In this case, we are doing a series of tests on ordinal data and the appropriate test is
a Komolgorov-Smirnov test (or KS test). The logic behind a KS test is not unlike a Chi2
test.
The first step is to sum the data across all trials for each experiment condition.
Then do the series of calculations as indicated in the following table
Response
Observed Frequency
0
0
1
6
2
8
3
9
4
13
Expected Frequency
7
7
7
7
7
Cumulative
Observed Frequency
Cumulative
Expected Frequency
Absolute value of
the difference
0
6
14
23
36
7
14
21
28
35
7
8
7
5
1
The critical KS statistic is the largest of the absolute values of the difference between the
cumulative observed and expected frequencies or Kmax = 8 . Look this value up in the
table provided at the end of the handout. In the example above, it will be for 5 classes of
data and 35 samples.
18 Summary of lab procedure
READ THE LAB!!!
Set up trials for male response
Petri dishes and glass rods
-control
-female pheromone extract
(each of 3 concentrations)
- bornyl acetate (a female
cockroach pheromone)
Score the response according to
the 4 point scale provided and
put the results on the board
Set up trials for female response
-put females into chambers
-1/2 hour acclimation (note position
and activity)
- 1/2 hour test period with stimulus
WEDNESDAY LAB
- Use live male & male
extract
Score the activity and
distribution results and put the
results on the board
THURSDAY LAB
- Use water & female
extract
19 REFERENCES
*August, C. J. (1971) The role of male and female pheromones in the mating behaviour
of Tenebrio molitor. J. Insect. Physiol. 17: 739-751.
Chapman, R.F. (1982) Exocrine glands, pheromones and defensive secretions. In: The
Insects, Structure and Function. 3rd Edition, Harvard University Press, pp. 851-885.
*Happ, G. M. (1970) Maturation and the response of male Tenebrio molitor to the
female sex
pheromone. Ann. Ent. Soc. Amer. 63: 1782.
*Happ, G. M and Wheeler, J. (1969). Bioassay, preliminary purification, and effect of
age, crowding, and mating on the release of Sex pheromone by female Tenebrio molitor.
Ann. of Entomological Soc. of America: 62:846-51.
*Tanaka, Y., H. Honda, K. Olsawa & I. Yammoto. (1986) A sex attractant of the yellow
meal worm,
Tenebrio molitor L. and its role in mating behaviour. J. Pesticide Sci. 11:
49-55.
*Tschinkel, Walter, C. Willson and H.A. Bern. (1967) Sex pheromone of the meal
worm beetle.
J. Exp. Zool. 167: 81-86.
Seabrook, W.D. and L.J. Dyer. (1983). Pheromones in insect control. In: Endocrinology
of Insects, pp. 673-686. Alan R. Liss. inc. NY.
*These papers are available on the course website at
http://www.mta.ca/~raiken/Courses/3401/Labs/labmanual.html
20 DATA SHEETS
1. MALE RESPONSE (Test 3 males in each condition!!!)
RESPONSE SCORE***
0
1
2
3
4
CONDITION
Control
Extract A
Extract B
Extract C
Bornyl acetate
***Record the number of males that reach each level of response for each condition.
2. FEMALE RESPONSE
A. ACCLIMATION PERIOD IN CHAMBER
POSITION OF FEMALES
NUMBER IN NEAR ZONE
15 MIN.
NUMBER IN FAR ZONE
30 MIN.
ACTIVITY OF FEMALES (during acclimation period)
15 MIN
FEMAL
E
1
2
3
4
5
6
30 MIN
Seconds(/3
0)
spent
moving
FEMAL
E
7
8
9
10
11
12
Seconds(/3
0)
spent
moving
FEMAL
E
1
2
3
4
5
6
Seconds(/3
0)
spent
moving
FEMAL
E
7
8
9
10
11
12
Seconds(/3
0)
spent
moving
21 2. FEMALE RESPONSE
B. TESTING PERIOD IN CHAMBER –
Male (WEDNESDAY) OR Clean glass rod (THURSDAY)
POSITION OF FEMALES
NUMBER IN NEAR ZONE
0
NUMBER IN FAR ZONE
3
6
9
12
15
18
21
24
27
30
ACTIVITY OF FEMALES
10 MIN
FEMALE Secs(/30) FEMALE Secs(/30)
moving
moving
1
7
2
8
3
9
4
10
5
11
6
12
30 MIN
FEMALE Secs(/30)
moving
1
2
3
4
5
6
FEMALE Secs(/30)
moving
7
8
9
10
11
12
20 MIN
FEMALE Secs(/30)
moving
1
2
3
4
5
6
FEMALE Secs(/30)
moving
7
8
9
10
11
12
22 B. TESTING PERIOD IN CHAMBER –
Male Extract (WEDNESDAY) OR Female Extract (THURSDAY)
POSITION OF FEMALES
NUMBER IN NEAR ZONE
0
NUMBER IN FAR ZONE
3
6
9
12
15
18
21
24
27
30
ACTIVITY OF FEMALES
10 MIN
FEMALE Secs(/30) FEMALE Secs(/30)
moving
moving
1
7
2
8
3
9
4
10
5
11
6
12
30 MIN
FEMALE Secs(/30)
moving
1
2
3
4
5
6
FEMALE Secs(/30)
moving
7
8
9
10
11
12
20 MIN
FEMALE Secs(/30)
moving
1
2
3
4
5
6
FEMALE Secs(/30)
moving
7
8
9
10
11
12
23 Assignment:
Every week you will have an assignment to complete. The lab
assignments are handed in the following week in lab. Make sure you are
doing the right assignment… the assignments for each week depend on
which group you are placed in (and therefore, which exercises you
completed). This is detailed below.
Group work and assignments:
Group A
Week 1
Exercise: Light and humidity preference in pillbugs AND snail/slug geotaxis and trail
following
Assignment: Write a formal results and discussion section for the work with the slugs
(the geotaxis work as well as trail following).
Week 2
Exercise: Chemoreception in flour beetles
Assignment: Write a formal abstract for the pheromone work with the flour beetles. This
means you must look at the data and include information on the results of the study and
the implications of this work. Abstracts tend to be tricky, read abstracts from other papers
to get an idea of content and layout.
Group B
Week 1
Exercise: Chemoreception in flour beetles
Assignment: Write an abstract for the pheromone work with the flour beetles. This
means you must look at the data and include information on the results of the study and
the implications work. Abstracts tend to be tricky, read abstracts from other papers to get
an idea of content and layout.
Week 2
Exercise: Light and humidity preference in pillbugs AND snail/slug geotaxis and trail
following
Assignment: Write a formal results and discussion section for the work with the slugs
(the geotaxis work as well as trail following).