Please see attachment named SP2021_Blackworm…. for instruction.

Please see other attachment for information. 

The lab report should be 4 pages long.

Short Lab Report Grading Checklist

This checklist may be used to double-check your short lab report before turning it in for grading. Point values for each item are given. The instructor reserves the right to make minor adjustments to the list before grading the report.

TITLE (5) – informative & concise

INTRODUCTION (15)

_____ There is only one Introduction section in the report. There are no other subtitles in this section. (3)

_____ The introduction provides the reader with enough information to know what the experiment is about and why it is being conducted. Sufficient information is provided that the reader can understand the rest of the lab report. The writer also explains why the hypotheses are being proposed, providing justification for them. (7)

_____ The hypothesis is stated and is clearly identified as the hypothesis that will be tested in the experiment. The hypothesis is appropriate for the observation being studied. The hypothesis is stated as a sentence that is either true or false. If the laboratory exercise has more than one hypothesis, they should all be stated. (5)

METHODS (15)

_____ This section contains only methods. There are no results or discussion in this section. (2)

_____ This section provides the reader with enough information that the reader could repeat the experiment. The writer uses their own words to write this section. (8)

_____ This section is written using whole sentences and paragraphs. Outline format, numbering, “cookbook” instructions, directions, or commands should not be used. (3)

_____ Avoid using the words I, he, she, or we. This section should be written in third person and there should not be references to other people. This section should be written in past tense. Example: “After the milk was added, three drops of rennin were added to tube #1.” (2)

RESULTS (25)

_____The following items are present in the results section: table of treatment, mean, standard deviation, standard error of the mean, and 95% confidence interval; graph of caffeine data with water control; graph of nicotine data with water control. (4)

_____ Only results are found in this section, nothing else. (2)

_____ This section is written using sentences and paragraphs. Any tables and figures in this section should be summarized using sentences and paragraphs in a narrative. (2)

_____ All of the results are included in this section. The results are written clearly and are easy to understand. (2)

_____ Descriptive statistics (e.g. mean, median, mode, standard deviation) are provided where appropriate. (2)

_____ At least one table is included. The table should be prepared by a computer. The table should be embedded within the Results section, not included on a separate page. (2)

_____ Each table has a caption that briefly describes the table. Example – Table 1. Reaction time 30 minutes after swimming 1000 meters at 8:00 AM. Be sure to provide enough information in the caption so that the reader can understand the table without reading the paper. Tables are labeled Table 1, Table 2, etc. At least one sentence in the Results section refers to the table(s). For example, “The reaction time decreased after swimming 1000 meters (Table 1).” (2)

_____ Figures have captions that briefly describe the figures. Be sure to provide enough information in the caption so that the reader can understand the figure without reading the paper. Graphs and other figures are labeled Figure 1, Figure 2, etc. At least one sentence in the Results section refers to the figure(s). For example, “The mean reaction times of the two groups differed (Figure 1).” (4)

_____ The axes of graphs are labeled correctly. The independent variable should be on the x-axis and the dependent variable is on the y-axis. Please see the notes on variables if you have any questions. (2)

_____ Tables and figures are placed in the Results section near where they are discussed, not at the end of the report. The tables are not printed on a separate page. Instead, they are inserted into the document. (1)

_____ The probability level (p-value) for statistical tests is stated. All of these statistics should be stated in sentence form. (2)

DISCUSSION (25)

_____ This section states whether you accept or reject the hypothesis or hypotheses. (5)

_____ If a hypothesis is rejected, an alternative hypothesis is given. (2)

_____ Acceptance or rejection of the hypothesis is supported by the data and statistical analysis. (5)

_____ Adequate interpretation of the data is provided. Unexpected results are discussed adequately. (8)

_____ Limitations to the experiment & dataset and possible future research are discussed adequately. (5)

REFERENCES (10)

_____ APA format is used for references. (3)

_____In-text citations are used. (2)

_____ Cited literature is appropriate. (5)

OTHER ITEMS (5)

_____ Point deduction if direct quotations were used. All citations must be reworded in the author’s own words. (2)

_____ The writer understands that hypotheses are not proven; they are accepted or rejected. (1)

_____ Complete sentences were used throughout. English is used correctly; grammatical errors are minimal. A spelling checker was used; there are no spelling errors or typographical errors. (2)

2

>S

1 8

_Thur2pm_

4

PR

–

Beats Pulse/Min

– Treatment Beats Pulse/Min

in rate

.3

12 0

12 13.3

12

4

.6

.3

.3

.3

.6

16

12 16 4
12

.3

8

20

16

1

20 2.7

16 20 4

7

28

12

.67

5.33

33

29

28 29 1

12

1

.6

16

12

17.3

27

30 15

mM)

12

28 9.3

16 32 16

21 25 4

21 28 7

4

1.33

5.33 1.33 -4
16 0

Nic ConC1 (0.05mM) 22 29 7
24

26 33 7

25

23 22 -1
27 21 -6

5.3

16 13.3

18.7 12

21 34 13

23 31 8
25 35 10

18.7 -4

22.7 25.3 2.6
17.3

Tea 21 25.3

19 25 6
19 30 11

20

25.3 12

18.7 12 -6.7

18.7 8

33.3 0

16 32 16

6 0

5.33

13.33 16

Tobacco 12 12 0
13.33 12

12

																																																																																																																																																																																																																																																																																																																																																																																																							Group 1		Before			Treatment		After		Change
				H2O													13								13.3							0
																																								12
1.3
			Caffeine ConC1 (1mM)											1			6
									14					1								7			2.7
																	16																21				5
							Group 2
			Caffeine ConC1 (1.0mM)										10	5.4
		17	5.3
			Caffeine conC 2 (3mM)												20	3			9										19
									28		36
							29		49
							Group 3
			Caffeine ConC 2 (3mM)		17.				33					1.33
7.3
			Caffeine ConC 3(10mM)					10.6	1				7.33
2				5.33				1	3.33
					18									24
							Group 4
						Water								32		-1
			34	-5
			Caffeine ConC3 (10mM)		2		9.3						17.3
4.6						30
	29.3
							Group 5
				Nic ConC1 (0.05mM)						27						23			-4
	42	–				15
-15
	Nic ConC2 (0.												25		37.3								25.3
									18.7
							Group 6
			Nic ConC2 (0.25mM)
														22										26
				Nic ConC3 (1.0mM)					13.33	-12
-16
						Group 7
					35										11
Nic ConC3 (1mM)						31		-6
						Group 8
	Cigarrette extract	2	2.6	-17.3
-2.7
	-6.7
					Tea
					Group 9
			Tobacco					22.7
	30.7	13.4
		4.3
					Group 10
		Cigarrette Extract							14.7	-5.3
-13.3
		Decaf				10.7
				33.3
			Group 11
Decaf			1		4.66				14.6
					9.33			14.66
2.67
-1.33
	10.66	-1.34

F17_Th2PM_4PS

Treatment Before After Change
Group 1

11 16 5

10

7.33

Group 2 Caffeine 3.0mM

7

20

11

Group 3 Caffeine 1.0mM 14 21 7

Extract

14.33

9.33

Group 4

7

2

Caffeine 10.0mM

Group 5 Caffeine 1.0mM 9

7

7.67

Caffeine 10.0mM

3

3

Group 6

9.67

7

13

12

7

Group 7

mM

21

15 19 4

Group 8

11 14.33 3.33

Nicotine 1.0mM 13 17 4

Group 9

Group 10

Group 11

Section 4PS
				Caffeine 1.0mM
				Caffeine 3.0mM		17.33
	5.6				14.33
				Caffeine 10.0mM		6.67	1			7.67
		Coffee	23.67
Decaf Tea		13.6		1	5.67
15.67		25.4	9.73
		16.6
					9.67			20.3			10.3
				Nicotine 0.25mM			22.6
			Nicotine 1.0mM		20.7	8.77
				Nicotine 0.05				12.67	8.33
Tobacco Extract
			Nicotine 0.05mM

F15_ThurNight

Group 1

H2O

33

66

8

Group 2

Caffeine ConC1 (1mM)

Caffeine conC 2 (3mM)

Group 3

Caffeine ConC 3(10mM) 10.7

Caffeine ConC 2 (3mM)

Group 4

Caffeine ConC1 (1.0mM) 11

Caffeine ConC3 (10mM) 10 24

Group 5

14.7 9.3

Cigarrette Extract

Group 6

Nic ConC2 (0.25mM) 18 8
Nic ConC3 (1.0mM)

7

Group 7

Nic ConC2 (0.25mM) 13 7.67
Nic ConC1 (0.05mM)

9.67

Group 8

Nic ConC3 (1.0mM)

6

Instant Coffee

6

2

Group 9

Cigarrette extract

23

Water

Group 10

Cigarrette Extract

7

Water

Pre-Treatment Beats Pulse/Min	Post- Treatment Beats Pulse/Min
	9.8		10.1
				Instant Coffee		5.666
	5.53		7.86
	5.32				11.66
2	4.5
	11.7		17.2
	16.666
		Nic ConC1 (0.25mM)
									15.3			11.3
	10.67
	12.33
14.	44
	13.5	1		8.2
		6.2		4.776
	6.223		6.776
	8.8
	7.76			8.1

SP16_4PR2

Group 1

Nicotine 0.05mM 15.3 13.3
Nicotine 0.25mM 15.3 14.7

Group 2

Nicotine 0.05mM 20.3

Nicotine 0.25mM

7

Group 3

26

22

Group 4

Nicotine 1mM 19 32
Caffeine 1mM 25 30

Group 5

9.33

12 11.66

Group 6

Caffeine 3mM

Caffeine 10mM 13.3 33.3
Instant Coffee 14 16

12 10.6

Group 8

10 26.6

Caffeine 10mM 7.3

Group 9

35 15.3

Nicotine 0.05 25 21.3

Group 10

Caffeine 3.0mM 4.5

4.3 4.66

	Pre-Treatment Pulse Pulse/Min		Post- Treatment Pulse/Min
		19.3
			18.6						21.3
	Nicotine 1mM		20.6
			Caffeine 1mM		31.3
			Caffeine 3mM		20.22
				Caffeine 10mM
	15.9				26.6
Group 7 Control
Spring Water
Green Tea
28.6
Tobbacco
9.5
Decafenated

F15_WedAM_

3EG2

3EG2 Water Caffeine 1.0mM Caffeine 3.0mM Caffeine 10.0mM Nicotine 0.05mM

Tea

Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp

Group 1 14.7

14 22.7

Group 2 19.3 17.3 17.3 26
Group 3 20.7

Group 4

24 20 24

Group 5 24 21.3

12.67

Group 6

26

Group 7

19 31
Group 8 18.6

21.33 22

Group 9

Group 10

13.33

Group 11 44

1

	Nicotine 0.25 mM		Nicotine 1.0 mM	Chewing Tobacco
																	Base																		Exp
	16.7
28.7
20.66
		16.67
	18.67	28.67	19.33
					21.33
57	71	22.89
	58		41
55.33	51	52
	17.99		9.1

F15_ThursPM_

4PS2

4PS2 Water Caffeine 1.0mM Caffeine 3.0mM Caffeine 10.0mM Nicotine 0.05mM Nicotine 0.25 mM Nicotine 1.0 mM Instant Coffee

Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp Base Exp

Group 1

21.33

9

Group 2 2.7 5.6 4.6 10.3
Group 3

16.6

Group 4 11.3 22 14

Group 5 10.1 25.3 13.6 21
Group 6 14 25 14.6 18
Group 7 13.3 15.3 12 15.3
Group 8

7.33

Group 9

14.6

26 22
Group 10 22 22

18 13
Group 11 13 11 14 10

Ciggarette
	24.4		20.8	28.89
14.9	20.4	29.5
43
	10.22
15.4

F15_TuesPM

Group 1 Pre-Treatment Pulse Pulse/Min Post- Treatment Pulse/Min

H2O 8 8.22
Instant Coffee

6

Group 2

Caffeine ConC1 (1mM)

Caffeine conC 2 (3mM)

Group 3

Caffeine ConC 3(10mM)

Caffeine ConC 2 (3mM)

Group 4

Caffeine ConC1 (1.0mM)

Caffeine ConC3 (10mM)

Group 5

Nic ConC1 (0.25mM)

9.55

8.1 13.5

Group 6

Nic ConC2 (0.25mM)

Nic ConC3 (1.0mM)

Group 7

6.2

Nic ConC1 (0.05mM)

	8.5		12.89
12.11	15.11
			9.55	13.44
11.67	13.77
8.66	11.22
9.45	13.67
8.45	13.11
10.45
	Nic ConC2 (.05mM)
15.667	12.5
	14.5	10.5
	Nic ConC2 (1.0mM)	6.89
	7.11	8.11

F15_WedNight

Group 1

H2O

8.5

H2O

9

Group 2

Caffeine ConC1 (1mM)

14.5

Caffeine conC 2 (3mM) 9.33 16.67

Group 3

Caffeine ConC 3(10mM) 10 20.8
Caffeine ConC 2 (3mM)

15

Group 4

Caffeine ConC1 (1.0mM) 9.55 13.33
Caffeine ConC3 (10mM) 9.55

Group 5

Nic ConC1 (0.25mM) 10.3 8.2
Nic ConC2 (.05mM) 9.1 9.8

Group 6

Nic ConC2 (0.25mM)

Nic ConC3 (1.0mM)

Group 7

Nic ConC2 (1.0mM) 7.33 10.33
Nic ConC1 (0.05mM)

7.11

Group 8

Tea

Pre-Treatment Pulses Pulses/Min	Post- Treatment Pulse Pulse/Min
9.443
10.367
9.86
11.53
15.87
8.87	17.6
	10.33	11.535
6.33
	9.43		9.77
Chewing Tobbaco		9.57		10.8

SP21_ShortReportData

Treatment

Treatment

Water 13.3 13.3 Water
12 12 Caffeine 1mM
12 13.3 Caffeine 3mM
33 32 Caffeine 10mM
34 29 Nicotine 0.05mM
28 29 Nicotine 0.25mM

Nicotine 1.0mM

6.223 6.776 Tea
7.76 8.1 Coffee
12 10.6 Decaf
14.7 16.7

24.4 21.33 Tobacco
Caffeine 1mM 12 16
14.6 17.3
16 21.3
10.6 16
12 16
12 17.3
11 16
14 21
9 16.67
5.53 7.86
11 16.666
22 31.3
Caffeine 3mM 20

28 36
29 49
16 17.33
17.3 20
16 20
10 17.33
5.67 20
5.32 11.66
11.7 17.2
9.33 20.22
15.9 26.6
Caffeine 10mM 10.67 28
12

18.67 24
12 29.3
14.6

12 29.3

6.67

15.67 25.4
9.67

10.7

10 24
12 11.66
Nicotine 0.05mM 27 23
42 27
30 15
22 29
24 35
26 33
12.67 21
11 14.33
12.33 9.67
15.3 13.3
20.3 19.3
25 21.3
Nicotine 0.25mM 12 37.3
18.7 28
16 32
21 25
21 28
22 26
9.67 22.67
18 8
13 7.67
15.3 14.7
18.67 21.3
16.67 12.67
Nicotine 1.0mM 13.33 1.33
5.33 1.33
16 0
31 25
23 22
27 21
12

13 17
10.67 7

6

26 20.6
19 32
Tea 21 34
23 31
25 35
21 25.3
19 25
19 30
10 26.6

19 31

18.6 21.33
9.43 9.77
Coffee 8 5.666
14 16

7.33 10.22
26 22

12.89

Decaf 10.7 18.7
33.3 33.3

16 32

14.66 14.66
9.33 14.66
13.33 16
4.3 4.66
Cigarette 22.6 5.3
16 13.3
18.7 12
20 14.7
25.3 12

18.7 12

15.3 11.3
6.223 4.776
8.8 7

18 13

Tobacco 22.7 18.7
22.7 25.3
17.3 30.7
12 12
13.33 12
12 10.66
35 15.3
21.33 22

58 41
17.99

9.57 10.8

Before Rate	After Rate	Change in Rate	[Stat1]	[Stat2]	[Stat3]	[Stat4]
9.833	10.166
	Cigarette
39
25.33
30.6
17.67
20.33
24.5
20.77
14.44
13.56	18.22
8.56
9.11

This article reprinted from:
Bohrer, K.E. 2006. Effects of drugs on pulsation rate of Lumbriculus variegatus

(blackworms). Pages 127-146, in Tested Studies for Laboratory Teaching, Volume 27
(M.A. O’Donnell, Editor). Proceedings of the 27th Workshop/Conference of the
Association for Biology Laboratory Education (ABLE), 383 pages.

Compilation copyright © 2006 by the Association for Biology Laboratory Education (ABLE)
ISBN 1-890444-09-X
All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or
transmitted, in any form or by any means, electronic, mechanical, photocopying, recording, or otherwise,
without the prior written permission of the copyright owner. Use solely at one’s own institution with no
intent for profit is excluded from the preceding copyright restriction, unless otherwise noted on the
copyright notice of the individual chapter in this volume. Proper credit to this publication must be
included in your laboratory outline for each use; a sample citation is given above. Upon obtaining
permission or with the “sole use at one’s own institution” exclusion, ABLE strongly encourages
individuals to use the exercises in this proceedings volume in their teaching program.

Although the laboratory exercises in this proceedings volume have been tested and due consideration has
been given to safety, individuals performing these exercises must assume all responsibilities for risk. The
Association for Biology Laboratory Education (ABLE) disclaims any liability with regards to safety in
connection with the use of the exercises in this volume.

The focus of ABLE is to improve the undergraduate
biology laboratory experience by promoting the
development and dissemination of interesting,
innovative, and reliable laboratory exercises.

Visit ABLE on the Web at:
http://www.ableweb.org

Association for Biology Laboratory Education (ABLE) 2005 Proceedings, Vol. 27:127-146

Effects of Drugs on Pulsation Rate of Lumbriculus
variegatus (Blackworms)

Kelly E. Bohrer

Department of Biology
University of Dayton

300 College Park
Dayton, OH 45469-2320

bohrerke@notes.udayton.edu

Abstract: In this investigative lab, students observe blackworm pulsation rate in normal conditions and
observe how pulsation rate is affected by drugs. This lab stresses the circulatory system, but can also be
used for homeostasis, behavior, toxicology, and nervous system labs. Part I guides the student through
blackworm handling procedures and initial observations of the blackworm’s behavior and circulatory
system. Part II is a student-led investigation in which the students design and run their own experiments
to test drug effects on pulsation rate. The students write their investigations as an informal report and
orally present their design, results, and conclusions.

Keywords: blackworms, Lumbriculus variegatus, pulsation rate, circulatory system, blood vessels,
student designed investigations

©2006 Kelly Bohrer

This major workshop paper is dedicated to and in memory of Dr. Charles Drewes.

Contents:
Introduction 128
Student Outline 130
Materials 138
Notes for Instructor 139
Acknowledgements 14

3

Literature Cited 143
About the Author 143
Appendix A: Recipes for Drug Solutions 144
Appendix B: Preparation Notes 145

128 ABLE 2005 Proceedings Vol. 27 Bohrer

Introduction

Background
Blackworms (Lumbriculus variegatus) are excellent organisms for studying the circulatory system

and the effects of drugs on this system for three main reasons: their skin is transparent making it easy to
observe pulsation rates, drugs quickly diffuse through the skin of blackworms thus providing immediate
effects, blackworms are easy to maintain in a laboratory. In blackworms, the dorsal blood vessel pumps
oxygenated blood from the posterior to the anterior end by muscular contractions in each segment. At
any time, several pulsation waves travel the length of the worm at a constant rate. Much like in humans,
the pulsation rate is regulated by the nervous and endocrine systems. Since many drugs affect these
systems (e.g. nicotine mimicking natural neurotransmitters), they can affect the rate of pulsation in
bloodworms. In this investigative lab, students observe blackworm pulsation rate in normal conditions
and observe how pulsation rate is affected by drugs.

In addition to the blackworm circulatory system, this lab stresses the following skills: scientific
process/inquiry, collaborative group work, critical thinking, verbal and written, data collection and
analysis, and working with live animals. Part I is designed to teach blackworm handling and viewing
procedures and to guide the student through initial observations of the blackworm’s behavior and
circulatory system. Part II is a student-led investigation in which the students develop their own
hypotheses and design and run their own experiments. The students write up their investigations as an
informal report and orally present their design, results, and conclusion at a later date.

In its current context, this lab exercise is completed in a two hour, non-major’s lab course. The lab
course is an introduction to biology that is meant to supplement the lecture course material. When this
lab exercise is performed, the students are learning about the human organ systems in lecture, including
the circulatory system. Prior to this lab exercise, the students have learned about the scientific process
and have designed a mini-investigation, including formulating an hypothesis, identifying a control,
identifying independent and dependent variables, analyzing results, and drawing conclusions.
Therefore, the lab content and process is not difficult for the students to understand; however, the
handling of the blackworms and the counting of the pulsation rate can be tricky. Therefore, it is
necessary that students are given sufficient practice with calculating pulsation rate before coming to lab
(by visiting the website indicated in the student outline) and given some time at the beginning of lab to
handle the worms (15-20 minutes). Other prerequisite knowledge and skills required for this lab include
microscope usage and evaluating outside sources of information on the internet.

This lab can easily be modified and/or expanded to a three hour and/or advanced biology lab (see
instructor notes). It can be adapted for lab exercises focused on homeostasis, toxicology, environmental
biology, behavior, or physiology. For example, students could calculate the Q10 of blackworms’
pulsation rate, test one or more physiological responses to external stimuli (pollution, acid rain, exercise,
salinity, etc.), observe regeneration of blackworm fragments, or explore acute and chronic exposure to a
toxicant.

There are many resources available for learning about blackworms and learning how to handle them
and experiment with them. In addition to the background information and reference publications in the
literature cited, you can also find a lot about blackworms on websites, especially Dr. C. Drewes website:
http://www.eeob.iastate.edu/faculty/DrewesC/htdocs. Additionally, teachers attending the 1996
Woodrow Wilson National Leadership Program have developed many similar blackworm lab activities,
which can be found on various websites.

Pulsation rate of blackworms 129

Lab Exercise Objectives

1. Identify blackworms.

2. Explain and identify key features and functions of the blackworm’s circulatory system.

3. Describe blood vessel pulsations of a blackworm.

4. Measure pulsation frequency and velocity.

5. Explain the effect of drugs on the circulatory system of blackworms.

6. Design and implement an investigation using blackworms.

7. Present results and conclusions both in writing and orally.

Timeline for Lab Activities
Culturing/buying worms Start culturing 2-4 weeks in advance
Cutting worms 24-48 hours in advance
Making solutions 24 hours in advance
Time needed for preparing lab ~5 hours if viewing slides have been

previously made

In-lab timing:
Introduction 10 minutes
Selecting and Handling worms 15 minutes
Determining Baseline Rate (Part I) 30 minutes
Designing Experiment (Part II) 20 minutes
Running Experiment 45 minutes

History of Blackworms in Biology Teaching Labs
Prior to 1996, Lumbriculus variegatus was well known among fish hobbyists. Thanks to Dr. Charlie

Drewes, the wonderful world of blackworms was introduced to biology teachers all over the nation by
Dr. Drewes’ Carolina Tips article in 1996 and by his guest appearance (as an instructor) at the 1996
Woodrow Wilson Institute at Princeton. Teachers from this institute have developed and shared many
blackworm related lab ideas, which has made this lab exercise possible. Additionally, much information
about culturing, handling, and viewing blackworms was gained through Dr. Drewes’ website and other
compositions.

130 ABLE 2005 Proceedings Vol. 27 Bohrer

Student Outline

Introduction

Purpose for This Lab
This lab activity serves three purposes: to introduce you to the circulatory system of blackworms, to

demonstrate the effects of drugs on the circulatory system of blackworms, and to provide additional
experience in designing and performing your own lab investigation. By the end of this lab, you will be
able to identify blackworms, explain and identify key features and functions of the blackworm’s
circulatory system, describe blood vessel pulsations of a blackworm, measure pulsation frequency and
velocity, explain the effect of drugs on the circulatory system of blackworms, design your own
investigation using blackworms, and present your results and conclusions both in writing and orally (due
at the end of the semester).

Function of a Circulatory System
A circulatory system is needed by any animal that is too large and/or complex to obtain essential

chemicals by the process of diffusion alone. Most importantly, a circulatory system quickly transports
nutrients, oxygen, and other important chemicals to all body cells. Circulatory systems have three
components: circulating fluid (blood or hemolymph), a heart or pulsating vessel which pumps the fluid,
and vessels through which the fluid travels. There are two types of circulatory systems, closed and
open. Open circulatory systems have vessels that are open at one end allowing hemolymph fluid to
flow out among the cells. Most mollusks and arthropods have an open circulatory system. In a closed
circulatory system, the fluid is called blood and this blood remains within the vessels as it rapidly
circulates the body. Vertebrates and annelids have a closed circulatory system. The pumping of blood
or hemolymph in a circulatory system is achieved by regular muscular contractions. The rates of these
contractions can be regulated either by hormones or by neurotransmitters released by nerve cells.

Lumbriculus variegatus
Blackworm is the common name for Lumbriculus variegatus, a freshwater oligochaete worm in the

phylum Annelida (earthworms and leeches are also in this phylum). Blackworms can be found
naturally in stagnant water along edges of marshes and ponds where they feed on small living and
decaying organisms. You can also find these worms at local tropical fish stores since they are great food
for pet fish. Blackworms are small worms, ranging from 4-6cm in length (~150 body segments with
head region containing 7-8 segments) in lab conditions, and up to 10cm in length in their natural
habitats.

Blackworms have several complex organ systems including a closed circulatory system, which
transports nutrients and oxygen; a complete digestive tract; and a nervous system, which includes a brain
and a nerve cord. Using their nervous system, the blackworm can respond very quickly to shadows,
touch, and vibrations by swimming, crawling, or performing a body reversal (rapidly coils and uncoils to
turn itself around). These worms obtain oxygen through their skin on their tail; hence the reason they
can often be found with their tails hanging out at the water surface. Unlike many other animals, sexual
reproduction is rare in blackworms; instead, it commonly multiplies by fragmentation and
regeneration. The worms will simply split into two or more sections, and each section will grow a new
head and/or tail. You may notice that some blackworms are darkly pigmented at one end compared to
the rest of the worm – the dark area is the original fragment (Drewes, 2003; Drewes 1996).

Pulsation rate of blackworms 131

Pulsation Rate of Lumbriculus variegatus
Today you will be observing the pulsation rate of blackworms. The blackworm has a large dorsal

blood vessel that is very easy to see using a microscope because the skin of the worm is transparent.
This dorsal blood vessel pumps oxygenated blood from the tail (which is usually kept towards the
surface of water) to the head of the worm by using rhythmic muscular contractions. The blood returns
to the posterior end of the tail via the ventral blood vessel, which is not pulsatory and is connected to the
dorsal blood vessel via small vessels in the first 1-18 body segments of the worm. In addition, to aid in
the pumping of the blood, most body segments have a pair of lateral, pulsatory vessels that do not
connect to the ventral blood vessel.

At any one time, you can see several pulsation waves along the length of the worm. Blood vessel
pulsation rate in blackworms is partially controlled by neurotransmitters that are secreted by nerve cells
(very similar to control of human heartbeats). The frequency (how many beats/waves per minute) and
the velocity (distance traveled per minute) of the pulsations can easily be calculated by observing the
pulse in the middle section of the worm (Lesiuk and Drewes, 1999). Because the rate of pulsation is
easily seen and calculated and some chemicals can easily diffuse through the worm’s thin skin, it is easy
to test the effects of exposure to different chemicals on the cardiovascular system of the blackworms.
This is what you will be doing for the second part of today’s lab. During the first part of today’s lab you
will be performing baseline observations of the behavior and pulsation rate of blacworms.

Safety Precautions, Disposal, and other Notes
1. Dispose of glass waste in the glass boxes
2. Handle organisms with care
3. Handle microscopes with care.
4. Report broken equipment, slides, etc. to the TA.
5. Making slides and cutting worms can result in minor wounds. Please take the necessary

precautions to avoid injury and report all cuts, however minor, to the TA.

Pre-Lab Assignment

1. Before lab begins, you will need to become familiar with blackworms and how to accurately measure
the dorsal blood vessel pulsation rate for the worms. Below is the URL for a website (Drewes, 2001)
that you should access before lab this week. This website provides a close up view of a blackworm
body segment (these are segmented worms) and shows you how blood pulsation occurs in a worm.
Read the directions and answer the questions for BOTH the posterior end of the worm and the mid-body
section of the worm.

http://www.eeob.iastate.edu/faculty/DrewesC/htdocs/INT-ANIMA-LvDBV-mid.htm

2. What types of chemical compounds affect the heart rate of humans? Perform an internet search to
find the names of at least two chemical compounds that affect the heart rate of humans. In what way
does the heart rate change when humans are exposed to these compounds (increase or decrease?) and
how does that change occur? Please remember to cite the name of any websites, books, articles, etc. that
you use.

132 ABLE 2005 Proceedings Vol. 27 Bohrer

Part One: Baseline Observations
In this part of the investigation, you will observe “normal” behavior and basal pulsation rate for

living blackworms. Make careful observations, sketch what you see, and record relevant data. Make
sure that both people get the chance to observe the worms’ pulsation rates using the microscope! At the
end of this initial investigation, you will be combining class data.

Important Notes
• The basal pulsation rate is generally greater at the tail end of the worm because many pulsations

starting at the tail end never make it all the way to the other end of the worm. Therefore, when
you observe the pulsation rate of the dorsal blood vessel, make sure to observe a mid-body
section of the worm and to always view the same segment throughout the entire investigation.

• Never use tap water with these worms! The chlorine in the tap water is toxic to the worms. Use
spring water and/or aged tap water for all parts of this experiment.

• Never use forceps or sharp objects to touch the worms – they are very fragile!
• Several factors can affect the behavior and the viewing of the worms = temperature, age, health,

direct light exposure, etc. Therefore, talk to your lab instructor if you have problems with a
particular worm.

Procedure

1. Fill both of your specimen bowls with spring water to a depth of approximately 2cm.

2. You will now select 5-10 worms that are equal in size. Avoid picking any worms that have
recently regenerated (worms that have a dark pigmented area and a lighter pigmented area). To
remove your worm from the water, you will need to use a plastic pipette. Gently suck up the
worm with a little bit of water and place into your specimen bowl.

3. You will also need to select 5-10 cut worms from the bowl at the TA desk. The anterior third
and the posterior third of the worm were cut off yesterday and placed in another bowl for
regeneration. You will be using the middle third of the worm for the second half of this lab since
this is easier to work with than a whole worm. The worms were cut yesterday to give the ends
time to heal for today’s lab. Before using these worm segments for the second part of today’s
lab, you will need to determine if the pulsation rate of the middle third is similar to the pulsation
rate of the mid-body segments of a whole worm. Why do you think we need to determine
this?

4. Obtain 5-10 middle third worm segments and place them in the second specimen bowl.

5. Watch the whole worms – what are they doing? How are they moving? Are they clumpled?
Swimming? Record in your notes what you observe. Can you identify the head end of your
worms? The head segments are generally darker, wider, and more blunt than the tail end. When
you observe these worms with the microscope, you should also be able to tell the difference
between the head and tail ends by how the blood vessel pulse moves (from tail to head).

6. Remove a whole worm from your bowl with a plastic pipette.

7. Place the worm into the trough on the well slide. Gently remove any excess water with a
chemwipe and place a coverslip over the worm. Wait a minute or two for the worm to adjust to
the trough (stop wiggling).

Pulsation rate of blackworms 133

8. Place the slide on the microscope and observe the worm at scanning power (4x) or using a
stereoscope.

*NOTE: Since intense light exposure can fry your worms and/or make them hyperactive, use a
low amount of light and avoid exposing your worms for long periods of time to the light.

9. Find a segment as close to the middle of the worm as possible. Count the number of pulsations
that pass through this point on the worm over 30 seconds. Multiply this by two to get rate per
minute. Repeat this procedure two more times. Then, find the average pulsation rate per minute
(record data in Table 1).

10. Place this worm into a weigh boat containing a small amount of spring water (just enough to
cover the worm). Label the weigh boat so that you can recall which worm is where.

11. Obtain another worm and have your partner run through #6-9 with this worm. This worm should
be placed in a different weigh boat. Record the data in Table 1.

12. Run through #6-9 using a third worm. Place this worm into its own weigh boat too. Record
your data in Table 1.

13. Now, run through #6-9 using three of the middle third segment worms. Record your data in
Table 1.

Table 1. Basal Pulsation Rate for Uncut and Cut Blackworms
Uncut Blackworms Average Pulsation Rate
1

2

3

Average Rate for Uncut:

Cut Blackworms
1

2
3

Average Rate for Cut:

Discussion

1. Put your results on the board. Your TA will calculate the average pulsation rate for both the cut
and uncut worms for the entire class. What is the class average for the cut worms? For the uncut
worms?

2. How did the class average pulsation rate for the cut worms compare to the class average for the
uncut worms?

134 ABLE 2005 Proceedings Vol. 27 Bohrer

3. Explain why the results are different or similar. What could have caused a difference if there is
one?

4. For data to be reliable, your data need to be accurate and reproducible. How have you achieved
this?

Part Two: Investigating the Effects of Drugs on Pulsation Rate in Lumbriculus variegatus
In this part of the lab, you will get the chance to design your own investigation. Certain chemicals

and drugs can greatly affect organ system function. Today, we will be looking at the effect of drugs on
the circulatory system of blackworms. Based on the research you performed for pre-lab and the
chemical compounds that your TA has available, you will design an investigation to see how pulsation
rate changes in response to exposing your worms to drugs. You have several options on how to design
this – you can investigate the effects of one or more drugs, investigate the effect of different
concentrations of a drug, investigate the effect of the length of exposure time to the drug, and/or you can
investigate the length of time it takes for the worms to recover from the effects of the drug.

Before beginning your investigation, please review information about setting up an investigation,
paying close attention to the necessary factors for a sound investigation (control, limiting variables, etc.).
Also, before you begin your procedures, talk to your TA about hints and suggestions for running this
type of investigation.

For your investigation, use the pre-cut worms (the middle body segments that you used in the
baseline observations earlier). If you need to use more worm segments, remember to first obtain a
baseline pulsation rate for each worm! Also, make sure to rinse your worms before placing them in the
trough and/or rinse your trough between observations so that you do not contaminate other worms.

Design Your Experiment
Before you begin, describe your experiment below and show your description to your TA. Do not

proceed with your experiment until your TA has given you the go-ahead.
• What would you like to investigate?

• What is your hypothesis?

• What is your dependent variable (what will you measure)?

• What is your independent variable?

o Why do you think this independent variable will affect the pulsation rate?

o How do you think this variable will affect the pulsation rate?

• What is your control? Be very specific!

• How will you include replications?

Pulsation rate of blackworms 135

• What results would support your hypothesis?

• Describe your methods:

• What materials will you need?

• What do you predict will happen?

Perform Your Experiment
As you carry out your experiment you will want to record your procedures, results (including table/s

to collect data and observations), and conclusions in a notebook. Be thorough and detailed as you
record your results. If you have problems, questions, and/or errors during the experiment, be sure to
write these down. Use the following information to guide you in writing your results and your
conclusions:

• Results – Describe your results in general. Do not explain why you got these results yet. Decide
how best to present your results – as a table and/or as a graph – and then complete your tables
and/or graphs before you interpret your results. You can use excel to design graphs.

• Conclusions –
o Look back at your hypothesis and look at your tables and graphs. Do your results support

or refute your hypothesis? Explain by using your data as evidence.
o Do your results match what you predicted above? Why or why not? Explain. If they are

not what you predicted, explain what may have occurred.
o If you had an opportunity to redo this experiment, how might you do it differently to

make it more convincing?
o Answer the summary questions below

Summary Questions
1. What was the reason for using more than one animal for each test? Did all animals respond in

the same way? Why or why not? What factors might influence individual response? What
implications does this have for the effects of drugs on humans?

2. Describe how the drug affected pulsation rate. Why do you think your results occurred?

3. Would your drug be classified as a depressant or a stimulant? Why?

4. What behavior characteristics did you note? Are they different than the behavior of the
unexposed worms?

136 ABLE 2005 Proceedings Vol. 27 Bohrer

Cleanup
1. Return all worm segments exposed to chemicals to the recovery bowl (do not dump chemicals

into this bowl –rinse your worms in spring water first).
2. Return all unexposed worm segments to the regeneration bowl.
3. Return all unexposed whole worms to the other bowl.
4. Dispose of chemical waste appropriately
5. Clean off all slides really well
6. Turn off your microscope, clean the lenses with microscope lens paper, and put away your

microscopes

Poster Presentation Information
Refer to Table 2 for grading information. You should present the sections of the poster on one

posterboard. You may use illustrations, pictures, drawings, etc. – be creative but don’t include irrelevant
information! Use a large font – something that can be read from 5 feet away. Single spacing is fine.
Label each part well (and in bold). All written parts should be in complete sentences and in paragraphs
(no bulleting).

Introduction:
The purpose of this section is to explain why you are performing this experiment and to provide

background information necessary to understand the framework of the experiment. Here you want to
describe the role of the cardiovascular system and discuss factors that can influence it (including “how”
and “why” these factors may change the pulsation rate). You also want to describe the organism we are
studying in lab and why we chose to use this organism. Then, you want to provide information about
the chemical you chose to test. Explain what is known about the chemical and then state what you
expected to see when you tested the chemical. Why did you expect this? The last paragraph typically
states your original question.

Materials and Methods:
This section should describe in moderate detail how the experiment was performed, and should

include the explanation of controls and the number of replicates performed.

Results:
This section is where the data is presented. Data should be presented as tables and graphs with

titles/brief explanations. No conclusions should be in this section.

Conclusions:
The conclusion should explain the results that you obtained and if your hypothesis was or was not

supported. This section should also include answers to any summary questions from the lab.

References:
Cite any outside information that you used when writing the introduction, material and methods,

and/or conclusions. See examples in book for correct format.

Pulsation rate of blackworms 137

Reflection Paper
Describe what you learned from this lab/process. Discuss what you liked about the lab and any ways

that this lab might be improved. 1-2 pages double-spaced.

Table 2. Grading Rubric for Investigation, Poster Presentation, and Reflection Paper
1 point 2 points 3 points 4 points Score

Prelab
Assignment

In complete in
more ways
than one

Partially incomplete,
no effort shown, no
references

Does not thoroughly discuss
#2, does not have accurate
answers for #1, and/or does not
reference websites

Completed on time,
correct information, #2
thoroughly discussed and
referenced

Question
Investigated

Not related to
topic and not
testable

Addresses too many
variables and/or not
related

Not in correct format, but is
testable and related

Directly related to prelab
research findings, testable,
correct format

Experimental
Design

Lacking 3 or
more of the
criteria for a
good
experiment

Lacking 2 of the
criteria for a good
experiment

Lacking one of the criteria for a
good experiment

Includes control, only one
experimental variable,
design directly answers
original question, other
variables kept constant

Poster
Introduction Question not

identified
and/or
summary
incomplete

Summary of
background
information is not
complete

Identifies question. Summary
of background information
complete, but not clear and/or
concise

Identifies question
investigated. Provides a
clear and concise
summary of necessary
info (see below)

Materials and
Methods

Not sequential,
most steps are
missing or
confusing

Some of the steps are
clear, most are
lacking detail and are
confusing

Most of the methods are
understandable, some lack
detail or are confusing

Clear and concise
summary of methods used
with adequate detail

Results Incomplete
information
including other
problems

Mostly complete
information, but
inaccuracies,
mislabeling, and
confusion

Information accurate. Labels
missing and/or information is
not clear

Tables and graphs
complete, accurate, well
labeled, and clear. Clearly
written summary of trends

Conclusions Presents an
illogical
explanation of
findings and
doesn’t address
original
question

Presents an illogical
explanation of
findings

Presents an explanation of
findings and addresses original
question, but is not clear and/or
complete

Presents a clear, complete,
and logical explanation of
findings, with evidence,
and addresses the original
question

References Missing
citations and
not in correct
format

Missing citations but
in correct format

Citations not in correct format Everything outside source
is cited, citations are in
correct format

Grammar Very frequent
grammar or
spelling errors

More than 2 errors Only one or two errors All grammar and spelling
are correct

Organization Disorganized,
incorrect
placement of
parts, not neat

Somewhat organized,
lacking flow,
incorrect placement
of parts

Mostly organized, some parts
are out of place or do not flow
well

Very well organized,
everything in correct
place, good transitions,
neat

Creativity Lacking
creativity

Creative, but the
creativity causes
design problems

Poster and question
investigated somewhat creative

Poster is creative and
question investigated is
unique and/or innovative

Reflection Paper Incomplete, no
depth, not
interesting

Somewhat
incomplete and
lacking depth

Complete, but lacking depth
and/or creativity

Complete, interesting,
creative, well thought out

138 ABLE 2005 Proceedings Vol. 27 Bohrer

Materials

Materials listed are for a class size of 20 students working in pairs. There are two student pairs at each
lab bench.

Culturing Blackworms
• Lumbriculus variegatus (need approximately 10-20 worms per group). These can be ordered from

Carolina Biological Supply (# CE-14-1720), Flinn Scientific, Inc (#LM1220), or can be purchased at a
local aquarium store. See appendix B for culturing instructions.

o “Starve” worms at least 2 weeks in advance of the lab
• Spring water or aged, dechlorinated water (let tap water sit in an open container for ~2 weeks)

o Worms are very sensitive to chlorine
• Small aquaria or buckets, large finger bowls, or 2 liter pop bottles for holding worms
• Brown paper towels
• Sinking fish food pellets

Prep Materials
• Single edge razor blades (new)
• Disposable petri dishes, Ward’s Biology (#19-7100)
• Filter paper, 90mm diameter, Ward’s Biology (#15-2815)
• Dissecting microscope
• Finger bowls to separate worms
• Standard Plastic pipets, Ward’s Biology (#18-2971)

Labroom supplies (front or back bench)
• Lumbriculus variegatus, uncut worms in one bowl, cut worms in another bowl
• Nicotine, caffeine, alcohol, and/or other drug solutions (labeled) (see Appendix A for recipes)
• Labeled plastic pipets
• Large finger bowl labeled “Rehab” and one labeled as “Regeneration”
• Graduated cylinders (1-ml and 10-ml)
• Latex gloves
• Spring water or aged, dechlorinated water
• Computer with internet access

Supplies at lab bench
• Parafilm trough slides or Tape well slides, 2 per group. (See Appendix B)
• Coverslips – heavy transparency or plastic preferable
• Chemwipes
• Weigh boats, 10 per pair, Ward’s Biology (#18-1453)
• Several standard plastic pipets
• Compound microscope and/or stereoscope, 1 per group
• Petri plates to raise worms away from light of stereoscope
• Widgets, 1 per group (See Appendix B)
• Eye droppers for solutions
• 100-ml beakers for solutions
• Stop watch – one per pair
• Cotton swabs
• Microrulers (See Appendix B)

Pulsation rate of blackworms 139

Notes for Instructors

Lab Design Information
The prelab assignment included in the student outline is a way for students to start making

observations about the blackworm’s circulatory system and about how drugs can affect heart rate.
Before this assignment is given, however, the students will need instruction on how to evaluate outside
sources of information on websites. They will also need to be reminded to cite all websites that they
reference. Coming to the class with the observations and background research complete, and an idea of
what type of drug they would like to test, saves time and prepares the students to make their hypotheses.
After Part I, the students formulate these hypotheses based on their observations, research, and supplies
available. The students will need guidance during this part so that their hypotheses are specific and
testable.

Before Part I begins, which is performed and discussed as a class, you may need to review basic
information about circulatory systems of annelids, “heart” rate, blackworms, designing a hypotheses and
making predictions, writing up lab results and conclusions, safety issues (see above), and the purpose for
the day’s activities. It will be important for the instructor to do background research before teaching this
lab to help guide students and work out issues that come up with using the worms. Refer to the
literature cited section of this paper for some of the best references on blackworms.

Purpose for Part I:
• To have baseline data (rates before treatment) to compare with experimental data.
• To practice handling and observing blackworms. Students will observe both behavior of the

worms and dorsal vessel pulsation.
Purpose for Part II:
• To design and run an investigation to test the effects of “x” on pulsation rate. Purpose is not

to kill the worms, but to determine the effects of sublethal concentrations of a drug on dorsal
vessel pulsation rate (and behavior).

Before having the students make their hypotheses for Part II, ask them what they found out about
drugs and the effects they have on heart rate. How do drugs affect heart rate? Is it dependent on
concentration and/or exposure time? How might exposure to several drugs change the effect? Can
blackworms recover from exposure? How long does it take? Can blackworms die from exposure?
Would acute exposure affect the pulse rate differently than chronic exposure? These are all good
questions for the students to consider in making their hypotheses for their experiment (Part II).

You can have the students proceed with their hypotheses and experiments in one of several ways:
• You pick which drug they test and how they test it.
• You pick which drug they test, but they choose what they want to test (concentration, type of

drug, exposure time, recovery time, etc.).
• Students pick which drug they test, but each group has to pick a different drug.
• Students pick which drug they test and it doesn’t matter if they possibly end up all picking the

same drug (you could have results about other drug effects available for them to see).
• Either of the last two option above, but you choose how they test it.

140 ABLE 2005 Proceedings Vol. 27 Bohrer

Possible Extension Activities:
• Calculating vessel diameter and pulsation velocity using microrulers (See Appendix B)
• Calculating Q10 of pulsation rate

o Q10 = rate at Temperature1 + 10 degrees/rate at Temperature1
• Chronic versus acute exposure to drugs
• Lethal and sublethal levels of drugs

o Students can determine the concentration of a particular drug that would evoke the
following responses:

 Low concentration = little to no response in pulsation rate
 Medium concentration = near maximum response
 High concentration = no increase response, but still sublethal

Viewing Blackworm pulsations
Look for worms that are healthy (wriggling), not recently regenerated (colorful), and “starved” (guts

are not dark). Use a plastic pipet to transfer the worms to their viewing slides (either the parafilm
troughs or the tape wells). Immediately after placing the worms on the slides, suck up extra water with a
fine tipped pipet. The water level should be even with the top of the well. Use tissue paper to soak up
extra water around the edges of the well. If the worm is not quite in the well or is wiggling out,
encourage it back in with the widget. Wait a minute or two for the worm to settle down. Then, place
the slide on the microscope and view using scanning power. Use a minimal amount of light so the worm
does not get overheated. Discern which way (to the left or to the right) that the pulse is moving so that
you can determine which end is the posterior end (remember, blood flows from posterior to anterior).
Pulsation rates may be higher at the posterior end of the worm because some of the dorsal vessel
contractions die out before reaching the anterior end of the worm. Therefore, it is important to monitor
the pulsation rates at the same location for each worm and for each time on the same worm.

Typical Data for Cut Vs. Uncut
Each number is an average for pulses/minute of three worms counted by each student group:

• Cut = 22, 13, 12, 16, 21 = 17 pulses/minute
• Uncut = 17, 13, 13, 13, 27 = 17 pulses/minute
• Cut = 18, 18.7, 14.6, 12, 17, 15 = 15.9 pulses/minute
• Uncut = 21.7, 10.6, 16, 14, 13.8 = 15.2 pulses/minute

How drugs affect the pulsation rate of Blackworms
Different chemicals (drugs) can have different effects on the pulsation rate of the dorsal blood

vessel. Their effect can occur by mimicking natural neurotransmitters that bind to “heart” receptors,
changing the propagation of action potentials, changing the amount of neurotransmitters released,
changing the amount and type of hormones that are released, blocking ion channels, or by directly
affecting muscular contractions. Most drugs enter the bloodstream of blackworms by diffusion through
the skin. Thus, they also wash out of the bloodstream once placed back in a dilute environment.

• Just like many other drugs taken recreationally, nicotine mimics a neurotransmitter that controls
pulsation rate. Nicotine is an acetylcholine agonist, so it increases the pulsation rate of the dorsal
blood vessel (depending on the concentration of nicotine).

Pulsation rate of blackworms 141

• In humans, caffeine inhibits the enzyme phosphodiesterase, thus allowing cAMP levels to go up
and ultimately increasing the heart rate. In blackworms, it is not clear if the increase in pulsation
rate due to caffeine is a direct or indirect effect.

• Alcohol acts as an ion channel blocker, thus decreasing the pulsation rate of the dorsal vessel.

Blackworm responses to solutions
• Response to caffeine – At low concentrations, worms may clump. As concentration increases,

worms become very active. They may curl up and stretch out at higher concentrations.
Pulsation rate will increase. Most worms recover within 15 minutes and all recover within one
day.

• Response to alcohol – Worms will become inactive as concentration increases and will be less
likely to clump. Worms may straighten out, with their ends curled, in higher concentrations.
They will not be able to swim as well. Their pulsation rate will decrease. Worms at the lower
concentrations will begin to recover within 15 minutes. Most all worms will recover within one
day.

• Response to nicotine – Worms become more active, but do not clump. At moderate doses, the
worms may be less active and twitch. In high doses (0.1 mM), paralysis (worm will be stretched
out and motionless) may occur. The pulsation rate may not show an increase at the lowest
concentration, but will increase at the middle and high concentrations. Most worms begin to
recover within 15 minutes and all recover within one day.

Tips for Instructors
• Suggestion for beginning the lab (to engage)-

o Have the students locate their pulse and ask them what they are sensing. Then, ask them
how they could change the rate of their pulse. Once they start bringing up many types of
drugs, ask them how drugs might affect the rate.

• When students first obtain their worms, have them place them in a weigh boat with just spring
water. Tell them to take some time making initial observations – which end is the tail end? How
do you know? What is the behavior of the worm? Does the worm swim? How? How is the
worm responding to its new environment? Etc. Lead a class discussion about their observations.

• To get the worm to stay in the well and to coax it into a good position for viewing, gently use the
widget or a piece of hair.

• Have one person view worm and count pulses while the other keeps track of time. Switch roles
often and make sure both students are relatively consistent when completing the baseline rate
count.

• Students can mix their own dilutions of the stock solution, or you can have dilutions already
made for them. They could also try other dilutions than just the ones recommended in the
recipes in Appendix A.

• Tell students to be careful not to transfer liquid from one container to the next as they move the
worms. Students also need to discern between pipets to reduce contamination.

• Wells need to be rinsed thoroughly with distilled water between worms

142 ABLE 2005 Proceedings Vol. 27 Bohrer

• Encourage students to thoroughly think about their control for their experiment. Moving worms
to and from dishes and slides could affect them; thus, it may be necessary to do the same for
control worms.

• You may want to check students’ procedures they have written down before letting them proceed
with their experiment. They may need a little guidance. Avoid telling them what to do; instead,
ask leading questions to help them develop a more sound experiment. Or, let them make
mistakes and talk about sources of errors and mistakes at the end of lab.

• If comparing pulsation rate of cut to uncut worms in Part I, have students put data on board.
• If students end up experimenting with more than just the few specimens they used for finding the

baseline rate in Part I, you will need to encourage them to find the baseline rate for all of the
other worms that they are going to use.

• Students may think that a very small change in pulsation rate is meaningless. Remind them that
even very small changes in an organisms’ body can have significant consequences. For example,
slight changes in body temperature, calcium levels, blood pressure, etc.

• Potential pitfall – Students will not notice a change or will have a myriad of problems resulting
in poor results. For these reasons, have the students also note changes in behavior so that, worse
comes to worse, they can write about behavior changes in their lab reports.

• If using statistics to analyze results, students will need to use at least 5 worms per treatment.
Students can use a paired difference t-test to compare before and after exposure.

• To save time, be strict about allotted times. Also, have data for the control group (worms that
are never treated but are transferred back and forth) already available.

After Lab Notes
• Rinse off worms well. Cut worms that were in treatments go into “Rehab” bowl (do not dump

treatment chemicals into this bowl!). Untreated cut worms can go into “Regeneration” bowl.
• Rinse off and dry viewing slides, weigh boats, and other containers.
• Throw away plastic pipets.
• Wipe microscope lenses with lens paper and turn off microscopes.

Pulsation rate of blackworms 143

Acknowledgements

I first want to thank Dr. Charles Drewes for all the help he offered me in preparing for this major
workshop. He was an incredible advice giver, a great support, and a friend – he will be missed. In
honor and in memory of him, I am dedicating this laboratory investigation write-up to him and the many
students that he has influenced throughout the years. I would like to thank both Charlie Drewes and the
teachers attending the 1996 Woodrow Wilson National Leadership Program for their work in
disseminating information about the use of blackworms in biology labs and for their lab investigation
ideas. I also would like to thank Dan Johnson from Wake Forest University for additional ideas and
feedback on this lab exercise and for Bunny for first encouraging me to use blackworms in the labs.

Literature Cited

Drewes, CD. 2003. A toxicology primer for student inquiry: Biological Smoke Detectors. The Kansas
School Naturalist, Emporia State University, 50(1):3-14.

Drewes, CD. 2001. Lumbriculus variegatus: A Biology Profile.
www.eeob.iastate.edu/faculty/drewesc/htdocs

Drewes, CD. 1996. Those wonderful worms. Carolina Tips, 59(3), 17-20.

Lesiuk, NM and Drewes, CD. 1999. Blackworms, blood vessel pulsations and drug effects. The
American Biology Teacher, 61(1), 48-53.

About the Author

Kelly Bohrer received a B.S. in Environmental Biology and an M.S. in Biology from The
University of Dayton, where she is currently the Biology Lab Coordinator. As such, she
coordinates the activities of 4 lab courses per semester; teaches biology labs, introductory
courses, and a graduate course on pedagogy for teaching assistants (TA’s); supervises TA’s and
prep assistants; and develops innovative lab curricula. Her research interests include wetland
ecology and laboratory pedagogy. She has recently received several grants to enhance laboratory
experiences for non-majors and pre-service teachers and to develop a university wide graduate
teaching assistant orientation.

144 ABLE 2005 Proceedings Vol. 27 Bohrer

Appendix A: Recipes for Drug Solutions

Make all solutions with spring water or aged, dechlorinated water. Use all solutions within 24 hours. Solutions
can be stored at room temperature. The dilutions are designed to give a small effect at the “low” concentration
and a more pronounced effect at the “medium” solutions. The “high” solutions should show that either the
threshold concentration has been reached or that the exposure response has reached maximum (but still sublethal).
Actual responses will vary depending on other factors.

Caffeine Stock Solution
• Use Vivarin tablets, NOT NoDoz!
• 200-mg caffeine/tablet
• Make a 5mM stock solution by crushing 2 caffeine tablets and add 412-ml of spring water (or aged

water). Dissolve tablets with stirring and heating if necessary.
• Use appropriate amounts of the stock solution to make 500-ml quantities of 0.1 (low), 1 (medium), and 5

mM (high) solutions.

Nicotine Stock Solution
• Cigarettes – regular length and strength, NOT menthol, 100’s, or ultralights
• 1.1mg nicotine/cigarette
• Make a 0.1mM stock solution by stirring the tobacco from 10 cigarettes in 680-ml of very warm spring

water for 20 minutes. Filter the solution. You will lose about 50-ml of the solution when filtering.
• Use appropriate amounts of the stock solution to make 500-ml quantities of 0.01 (low), 0.05 (medium),

and 0.

1 mM (high) solutions.

Alcohol Stock Solution
• Vodka = 40% alcohol
• 1 mM alcohol = 2.6%
• Mix 32.5-ml of vodka and 467.5-ml of spring water to make the stock solution.
• Use appropriate amounts of the stock solution to make 500-ml quantities of 0.1 (low), 0.5 (medium), and

1 mM (high) solutions.

Other Possible Drug/Toxicant Solutions
• Diet pills, cold medicine, Tylenol, acetylcholine, epinephrine, lidocaine, glucose, sugar substitutes, saline

solution, detergents, pesticides, etc.
• Crush and dissolve tablets in spring water. Make a high, medium, and low solution. Groups could work

to find the concentrations that lead to little response and maximum response.

Pulsation rate of blackworms 145

Appendix B: Preparation Notes

Culturing Blackworms
Fill a bucket, small aquarium, or large finger bowl with 2-3 inches of aged, dechlorinated water (or spring

water). Add healthy worms (about 100) to the water and then layer the water with several small pieces of brown
paper towel. Every week add one to two (depending on size of aquarium) pellets of sinking fish food. Do not
overfeed! As water evaporates, add spring water to the original level. When the water begins to appear cloudy
and/or starts to stink, slowly pour off as much of the water as possible without losing the paper towel pieces or the
worms. Rinse the worms and paper towel once (with aged water) and then refill the aquarium (to 2-3 inches) with
fresh water and a few new pieces of brown paper towel. If you are not using the worms for a while, split the
culture or feed some of the extra worms to fish (the culture should double every 2-3 weeks and more quickly with
slight agitation). This culture should live for a long time following these procedures.

Handling Blackworms
Worms are best handled by sucking them up with a plastic disposable pipet. Blow out the air in a pipet, place

the pipet at a 45 degree angle, lower it to the bottom, and quickly suck up 1 or 2 worms at their head ends. If you
dismember any of the worms in the process, just leave the pieces in the culture to regenerate.

Using Cut Blackworms
Blackworms that have been fragmented tend to move around less. Typically, the pulsation rate of a newly

fragmented worm is close to the pulsation rate of a whole worm. At UD, we have the students actually verify this
before “choosing” to use cut worms for their experiment. Each student group measures the pulsation rate of three
cut and three whole worms (in approximately the same region of the worms), and then pool their data with the rest
of the class. Then, as a class, we can decide if the pulsation rates are close or not. To save time, it may be best to
either tell them that this is so or to have data available for whole worms and have the students see for themselves.
If you do choose to use cut worms, these worms should be cut at least one day in advance so that the ends are
healed. Select whole worms that are healthy and full-sized and cut them into thirds by placing them on a piece of
saturated filter paper in a petri dish and cutting them with a clean, sharp razor blade. Keep the middle segments
for the experiments and put the other two segments back into your culture so they can regenerate. Keep the
middle segment worms separate from the whole worms and place both into separate bowls with fresh spring water
for the lab exercise (label the bowls as “cut” and “whole”).

Making “Widgets” (for moving worms)
The following directions for making widgets is adapted from “A Toolbox for Working With Living
Invertebrates,” by Dr. Charlie Drewes. This article can be found in ABLE’s 2004 proceedings.
1. Materials: applicator stick (handle of a probe works well), rubber band, scissors, and tape.
2. Cut, at an angle, a piece of rubber band that is one inch long.
3. Attach the rubber band to one end of the applicator stick with tape, leaving _ inch of rubber band beyond the

end of the stick.

Making Viewing Slides
The procedure for making tape well slides is also presented in the article listed above. You can also find
directions, and pictures, for both widgets and slides at
http://www.eeob.iastate.edu/faculty/DrewesC/htdocs/ (scroll down to “Gadgets & Technical Information”)

Tape Well Slides:
1. Materials: clear plastic tape (Scotch Colored Plastic Tape, Clear, 0.75” X 125”); forceps; single edge

razor blade; heavy scissors; heavy-duty, flexible clear plastic (or glass microscope slides); pen; ruler
2. Using a pen and a ruler, mark off desired size slides on the plastic sheet.

146 ABLE 2005 Proceedings Vol. 27 Bohrer

3. Place a long strip of tape over the plastic sheet, ensuring that there are no bubbles
4. Add multiple layers of tape (4-5).
5. Using a ruler and razor blade, make vertical cuts to define the well sizes for holding the blackworms

(3-mm deep and wide and 4-cm long)
6. Using a forcep, carefully lift the tape layers covering the desired well.
7. Cut out the “slides” from the plastic sheet.
8. On another piece of clear plastic, mark off and cut out rectangles to act as cover slips for your slides

(make them a little smaller than your slides).

Parafilm Trough Slides:
1. Materials: single edge razor blade, parafilm, glass microscope slides, ruler, metal surfaced hot plate,

glass plate (~same size as hot plate), forceps, glove for hot items
2. Put hot plate on low.
3. Lay out glass slides, side by side, on glass plate to about 6” long.
4. Cut out pieces of parafilm that are 4” X 6”.
5. Place several layers of parafilm (6-8) on the glass slides and press down on the parafilm to make sure

the sheets stick to each other and the glass slides.
6. Put glass plate on hot plate and let it warm up for ~5-10 minutes. During this time, use parafilm

backing to press the softened parafilm against the slides. Try to remove all air bubbles and make sure
everything is sticking together.

7. Once the parafilm just begins to get clear and soft, remove the glass sheet (with gloves) and carefully
place it on the counter.

8. Using a ruler and razor, make cuts in parafilm to define the well sizes for holding the blackworms (3-
mm deep and wide and 4-cm long). Make long wells for whole worms and short wells for cut worms.
You can put two short wells and one long well on each slide.

9. Use forceps to remove the parafilm from the cut wells.
10. If part of the parafilm lifts during this time, simply reheat the slide on the hot plate and press down on

the parafilm.

Making Microrulers
• Visit www.eeob.iastate.edu/faculty/DrewesC/htdocs/microruler-links.htm

Safety Issues
• Clearly label the contents and concentrations of all chemical solutions, including stock solution and

dilutions (remind students to do this).
• Read the Material Safety and Data Sheets (MSDS) for chemicals being used. For chemicals that are

health hazards, including nicotine, wear gloves and minimize contact.
• Properly dispose of all solutions and all materials exposed to solutions.
• Scrub and clean all glassware and wipe down all benches with ethanol at the end of lab.
• If accidentally cut, wash the area thoroughly.
• Handle microscopes appropriately. Use lens paper to clean lenses before and after use.

38 Volume 43(2) December 2017 Ryan and Elwess

A New Approach in Examining the Influence of Drugs on Pulsation Rates in
Blackworms (Lumbriculus variegatus).

Amy B. Ryan and Nancy L. Elwess*

Department of Biological Sciences, State University of New York at Plattsburgh, Plattsburgh, NY 12901

*Corresponding Author: elwessnl@plattsburgh.edu

Abstract: This investigative laboratory activity engages students in observing, recording,
graphing and analyzing pulsation rates in a commonly used laboratory organism, blackworms.
This activity stresses how various drugs can impact the pulsation rate in blackworms at varying
concentrations. In addition, we have incorporated two new ways to view the blackworms under
the microscope.

Key words: Blackworms, pulsation rate, Lumbriculus variegatus, blood vessels, capillary tubes

INTRODUCTION
Lumbriculus variegatus, or blackworms,
are freshwater oligochaetes in phylum
Annelida. They are an excellent organism
for studying the regeneration of body parts,
regulation of reflex activities,
bioaccumulation and toxicity of
environmental pollutants, and regulation of
blood vessel pulsations (Drewes & Fourtner,
1990; Veltz et al., 1996; Bohrer, 2006;
Fillafer & Schneider, 2013).
Like other annelids, blackworms have a
closed circulatory system (Fig. 1).
Blackworm blood is red, due to a
hemoglobin-like pigment called
erythrocruorin dissolved in the blood plasma
(Jamieson, 1981). Two major blood vessels,
one dorsal and one ventral, extend the length
of the blackworm. Pulsations along the
dorsal blood vessel (DBV) propel blood
through the circulatory system. Because the
body wall of the blackworm is transparent, it
is possible to visualize the pulsation of the
DBV using light microscopy (Lesiuk &
Drewes, 1999). As in humans, the pulsation
rate is controlled by the nervous and
endocrine systems. Many drugs affect these
systems and can have an immediate impact
on the pulsation rate, based in part on how
quickly they can diffuse through the
blackworm’s skin. Due to their simple body

plan and ease with which they can be treated
with compounds and the subsequent
pulsation rate measured, black worms are an
excellent model organism for the lab activity
described below.

Fig.1. A lateral cross section image of a closed circulatory
system found in blackworms. Included in this image are
major blood vessels and the internal anatomy. The
pulsation rates can be determined by counting the pulsation
waves at one location on the dorsal blood vessel. Image
created by Sophie Kim.

Influence of Drugs on Pulsation Rates Bioscene 39

This investigative laboratory, designed for
a college freshman general biology course,
takes a fresh look at a standard blackworm
laboratory activity that was first published in
1999 (Lesiuk & Drewes) and again in 2006
(Bohrer, 2006). Lesiuk and Drewes describe
how blackworms can be used as a model
system to demonstrate the effects of nicotine
and caffeine on blood vessel pulsation and
explain how to make the blackworm
viewing chambers. Bohrer provides a more
in-depth background as to how the Lesiuk
and Drewes preparation can be incorporated
into the curriculum, by including timelines,
materials, methods, and a suggested grading
rubric. We have developed two new
approaches for viewing the blackworms
under the microscope (Worm Viewing
Chambers under Procedures) and have
added three drugs to those from the original
publications. This activity was done over
three weeks. Week 1 (Wk 1) instructed the
students on blackworm handling and
behavior, as well as determining the
pulsation rate under control and
experimental conditions (varying caffeine
concentration). Wk1 served as the practice
week for procedures done in week 2. Week
2 (Wk 2) was inquiry, with each pair of
students exposing their worms to one of
three new treatments (cinnamon, celery seed
extract, and valerian root). Wk 2 served as
the application week. Both Wk 1 and Wk 2
stressed laboratory skills, including
scientific inquiry, data collection, and
microscopy. Scientific inquiry included the
students becoming skilled at stating a
hypothesis (Wk 1), determining the controls
(Wks 1 & 2), calculating drug
concentrations (Wks 1 & 2), determining
results (Wks 1 & 2), and in Week 3,
graphing and data analysis used in reaching
their conclusions. The laboratory protocol
can be easily altered for an advanced
biology activity by involving more
chemicals, examining reaction to a variety
of stimuli (i.e. temperature), and
regeneration rates.

PROCEDURES
Blackworm Care
Blackworms were obtained from
Carolina Biological and kept in a small tank
containing approximately 2 inches of
aerated spring water and strips of brown
paper towel. They were fed fish food flakes
every two weeks. When not in use, the tank
was placed in the dark.
Worm Viewing Chambers
We came up with two very distinct, but
easy to use viewing chambers for the
blackworms. In the original publication
(Lesiuk & Drewes, 1999), a viewing
chamber was made by using six layers of
Parafilm bonded onto a microscope slide
using heat. The trough, which held the
blackworm, was made by cutting the
Parafilm with a razor blade. Using a 3D
printer, we designed a plastic slide with the
trough embedded, and simply glued this
onto a microscope slide (Figs. 2 & 3). A
Makerbot Replicator 2 and the Solidworks
3D CAD software program was used to
generate the viewing slide. The dimensions
for the viewing slide was 7.5 cm long x 2.5
cm wide x 0.2 cm high, and the
trough slot embedded within the slide was 4
cm long x 0.2 cm wide x 0.2 cm high. Once
these dimensions were entered into the
software program, it was exported into

Fig. 2. At the top is the traditional parafilm made trough as
described in Lesiuk and Drewes (1999). In the middle is
our 3D made trough and at the bottom are 100 µl capillary
tubes. All can be used for viewing blackworms under the
microscope. Photo taken by N.L. Elwess.

40 Volume 43(2) December 2017 Ryan and Elwess

Fig.3. On the microscope slide is a blackworm in a 3D
printer made trough. Photo taken by N.L. Elwess.

Makerbot’s software. Our second viewing
chamber was a 100 µl glass capillary tube
(Figs. 2 & 4) with a diameter of 1.35 mm. A
worm can be easily transferred with a plastic
Pasteur pipette into the capillary tube (Fig.
4). The capillary tube was then placed
directly onto the microscope stage, where it
could be easily rotated to obtain the best
view of the DBV.
Prior to Starting this Laboratory Activity
The week prior to starting this laboratory
activity, students were given a pre-lab
assignment that was due the first day of this
lab. Included in this pre-lab assignment were
questions students needed to answer based
on their reading of the introductory material
in their laboratory manual. These questions

Fig. 4. A. Image of blackworm as seen in capillary tube
under no magnification B. Image of blackworm as seen in
capillary tube under 40X magnification Scale bar
represents 1000 mm. Photos taken by N.l. Elwess.

covered their knowledge of why
blackworms would make a good model
organism for determining pulsation rates,
parts of a dissecting microscope, and asked
the students to provide a hypothesis for the
effect of caffeine on blood vessel pulsation.
The laboratory activity prior to this
experiment introduced students to the nature
of science, scientific inquiry, and basic
experimental design terminology. This
activity we felt was a good follow up to
build on their understanding of those
concepts. This article focuses on
improvements made for the viewing of the
blackworms and some suggestions for new
experimental approaches to those previously
published (Lesiuk & Drewes, 1999; Bohrer,
2006). Bohrer (2006) does an excellent job
describing the lab set-up, student learning
objectives, materials needed, and a
suggested grading rubric.
Week 1
During Wk 1, the students became
familiar with handling the blackworms and
determining the basal pulsation rate. Each
pair of students selected three blackworms
from the tank and transferred them, using a
plastic Pasteur pipette, into three separate
petri dishes. Once students became
comfortable with handling the blackworms,
they started counting the DBV pulsation rate
for each blackworm; each pulsation rate was
done in triplicate. Students viewed the
blackworms under a dissecting microscope
at 20x magnification. Four different mM
concentrations of caffeine were used (0.1, 1,
5, and 10). Working in pairs, students
counted and recorded the basal pulse rate for
each worm, this served as a control, and then
tested one of the four caffeine
concentrations. Students made 30 mL of the
test solution by diluting a 20mM caffeine
stock solution. Each worm was placed in
10mL of test solution. A blackworm was
exposed to the test solution for 15 minutes
prior to counting the pulsation rate. Students
recorded and shared their data using Table 1.

Influence of Drugs on Pulsation Rates Bioscene 41

Week 2
Three different compounds were used to
determine if they had an influence on
pulsation rate. All lab groups repeated the
same approach as in Wk 1, but in Wk 2
instead of testing caffeine, they tested two
concentrations (0.1 mg/mL and 1.0 mg/mL)
of Cinnamon, Celery seed extract, or
Valerian root. Three worms were observed
at each concentration and each blackworm
was placed in a single petri dish. Cinnamon,
a spice widely used in traditional medicine,
has been linked to a reduction in
cardiovascular disease due to its effects on
excitability (Alvarez-Collazo et al., 2014).
Celery seed extract, which has been used in

the Eastern world for thousands of years, is
a diuretic used in the treatment of high blood
pressure (Moghadam, et al., 2013). Valerian
root is an herbal supplement used to reduce
blood pressure and prevent arrhythmias
(Chen et al., 2015). Students calculated the
appropriate dilution needed to make 30 mL
of test solution from the commercially
purchased stock solutions of cinnamon
(905.7 mg/mL), celery root extract (870
mg/mL), and valerian root (1000mg/mL).
Students recorded and shared their data
using Table 2.
Week 3
Each student generated graphs of the
average pulsation rates for each experiment

42 Volume 43(2) December 2017 Ryan and Elwess

using Microsoft excel (Figs. 5-7). Using the
graphed data, the students determined if
their initial hypotheses were supported or
refuted and then wrote their conclusions.

Fig. 5. An example of a student generated graph showing
the results of the effect of caffeine treatment on pulsation
rates in blackworms. There was a significant difference at
0.1 mM, 1.0 mM, and 5.0 mM concentrations of caffeine.
Each of those experimental conditions had a high pulsation
rate over the control.

Fig. 6. An example of a student generated graph showing
the results of the effect of celery extract on pulsation rates
in blackworms. At both concentrations (0.1 mg/mL, 1.0
mg/mL) there was a significant increase in pulsation rates
over that of the control.

Fig. 7. An example of a student generated graph showing
the results of the effect of cinnamon on pulsation rates in
blackworms. At the 1.0 mg/mL there was a significant
increase in pulsation rates over that of the control, while the
0.1 mg/mL concentration did not show any difference in
pulsation rates.

RESULTS AND DISCUSSION
Overall, the updated worm viewing
chambers used in this lab allowed students
to better manipulate the worms under the
microscope for optimal viewing of vessel
pulsation. The 3D generated viewing
chamber worked well for viewing larger
blackworms, and the capillary tubes worked
well with any size blackworm. The second
week of the activity, where a novel
compound was tested for its effect on
pulsation, provided students the opportunity
to compare treatment groups and perform
statistical analyses. The inclusion of novel
compounds also required the students to
perform literature searches for candidate
molecules and gave them the opportunity to
develop testable hypotheses and make
predictions based on their findings. As an
experimental modification, students could
also research and select the novel compound
that they will assess, and design their own
experiments, thereby introducing another
component of scientific inquiry. They could
also assess the effect of their compound on
additional parameters, including blackworm
behavior, reflex activity, or regeneration.
ACKNOWLEDGEMENTS
We thank Dr. Michael Walters, Assistant
Professor in the Physics Department at the
State University of New York at
Plattsburgh, for the design and making of
the trough slides via a 3D printer. We would
also like to thank Sophie Kim for the
creation of Figure 1.
REFERENCES
ALVAREZ-COLLAZO, J., ALONSO-
CARBAJO, L., LÓPEZ-MEDINA, A. I.,
ALPIZAR, Y. A., TAJADA, S., NILIU, B.,
AND J.L. ALVAREZ. 2014.
Cinnamaldehyde inhibits L-type calcium
channels in mouse ventricular
cardiomyocytes and vascular smooth muscle
cells. Pflugers Archiv: European Journal of
Physiology, 466(11), 2089–2099.
https://doi.org/10.1007/s00424-014-1472-8

Influence of Drugs on Pulsation Rates Bioscene 43

BOHRER, K.E, 2006. Effects of drugs on
pulsation rate of Lumbriculus variegatus.
Tested Studies for Laboratory Teaching,
27:127-146.
http://www.ableweb.org/volumes/vol-
27/07_Bohrer

CHEN, H.-W., WEI, B.-J., HE, X.-H., LIU,
Y., AND J. WANG. 2015. Chemical
Components and Cardiovascular Activities
of Valeriana spp. Evidence-Based
Complementary and Alternative Medicine:
eCAM, 2015, 947619.
https://doi.org/10.1155/2015/947619
DREWES, C.D. AND C.R. FOURTNER.
1990. Morphallaxis in an aquatic
oligochaete, Lumbriculus variegatus:
Organization of escape reflexes in
regenerating body fragments.
Developmental Biology 138:94-103.
FILLAFER, C. AND M.F. SCHMEIDER.
2013. On the temperature behavior of pulse
propagation and relaxation in worms,
nerves, and gels. PLOS One, June 21, 2013,
http://dx.doi.org/10.1371/journal.pone.0066
773
JAMIESON, B. G. M. 1981. The
ultrastructure of the oligochaeta. Academic
Press.

LESIUK, N. M., AND C.D. DREWES.
1999. Blackworms, Blood Vessel Pulsations
& Drug Effects. The American Biology
Teacher, 61(1), 48–53.
https://doi.org/10.2307/4450609
MOGHADAM, M. H., IMENSHAHID, M.,
AND S.A. MOHAHERI. 2013.
Antihypertensive effect of celery seed on rat
blood pressure in chronic administration.
Journal of Medicinal Food, 16(6), 558–563.
https://doi.org/10.1089/jmf.2012.2664
VELTZ, I., ARSAC, F., BIAGIANTI-
RISBOUR, S., HAVETS, F.
LECHENAULT, H., AND G. VERNET.
1996. Effects of platinum on Lumbriculus
variegatus: acute toxicity and
bioaccumulation. Arch. Environ. Contam.
Toxicol. 31:63-67.