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TEACHING IN THE LABORATORY
Department of Physiology, Faculty of Medicine and Dentistry, University of Western Ontario, London, Ontario, Canada N6A 5C1
Abstract
To address a growing need to make research trainees in physiology comfortable with the tools of molecular biology, we have developed a laboratory-intensive course designed for graduate students. This course is offered to a small group of students over a three-week period and is organized such that comprehensive background lectures are coupled with extensive hands-on experience. The course is divided into seven modules, each organized by a faculty member who has particular expertise in the area covered by that module. The modules focus on basic methods such as cDNA subcloning, sequencing, gene transfer, polymerase chain reaction, and protein and RNA expression analysis. Each module begins with a lecture that introduces the technique in detail by providing a historical perspective, describing both the uses and limitations of that technique, and comparing the method with others that yield similar information. Most of the lectures are followed by a laboratory session during which students follow protocols that were carefully designed to avoid pitfalls. Throughout these laboratory sessions, students are given an appreciation of the importance of proper technique and accuracy. Communication among the students, faculty, and the assistant coordinator is focused on when and why each procedure would be used, the importance of each step in the procedure, and approaches to troubleshooting. The course ends with an exam that is designed to test the students’ general understanding of each module and their ability to apply the various techniques to physiological questions.
Key words: graduate education; hands-on approaches
The past decade has seen an explosion in molecular biological techniques designed to answer many questions that previously were unapproachable. The human and mouse genome projects will bring even wider use of molecular techniques, particularly in the emerging field of physiological genomics. With this explosion has come the challenge to provide trainees with the background and expertise required to use molecular tools to address their research problems. This is especially true for graduate students who, during their undergraduate education, may have had limited exposure to molecular biology or little hands-on experience with molecular techniques. To address this concern, we have designed a course to teach beginning graduate students in the Department of Physiology at the University of Western Ontario the basic techniques for manipulating DNA and RNA, and for studying the expression of specific mRNAs and proteins in organs and cells. Practical, hands-on laboratory experience, coupled with lectures that explain the theoretical background for the techniques and their application in physiological research, make this course a valuable introduction to molecular biology.
The three-week intensive course is organized into modules, each one organized and supervised by a faculty member. In this respect, the course capitalizes on the expertise of various faculty members within the department and provides students with a multidisciplinary view of how molecular biology can be used to answer a wide range of questions. Students are also exposed to the research interests and expertise of participating faculty. Having a team of organizers in a hands-on course such as this also serves to maximize experimental success, given that each protocol will have been optimized in the laboratory of the investigator responsible for that module.
The outline provided in this report is the result of numerous improvements and is based on strategies proven to be effective, as this course has been offered for four consecutive years in our department. In the first year, the course was offered as a "trial run," with no formal evaluation of participants. Typically, the course is offered to a maximum of eight students who are enrolled in the graduate physiology program. When space is available, graduate students from other departments as well as technicians or postdocs within the Department of Physiology are encouraged to enroll. The students work in pairs, both to facilitate collegial interactions and to conserve reagents.
The feedback from this course has been extremely positive, with students commenting on their enjoyment of the course and the effectiveness of the curriculum in providing them with a very clear understanding of molecular techniques. In addition, the students’ supervisors are impressed by the benefits that training such as this offers to their own research programs. Although they do not finish the course as experts, the students do take away sufficient knowledge to be able to apply molecular techniques in their own research. In addition, when combined with the lectures, the labs offer background knowledge that is invaluable for interpreting scientific literature and data presented in seminars, two major components of their training. Given the success of this course and the extensive effort that has gone into its organization, we have written this report to assist other departments in adding a similar course to their program.
COURSE OUTLINE
The course is organized into seven modules, each aimed at introducing a specific set of tools in molecular biology. These modules are as follows:
Module 1: DNA: properties and common molecular biological terminology; cDNA subcloning
Module 2: gene transfer and protein expression analysis
Module 3: RNA expression analysis
Module 4: reverse-transcriptase polymerase chain reaction (RT-PCR)
Module 5: in situ hybridization with immunochemical detection of hybridized probe
Module 6: DNA sequencing and use of sequence databases (lecture and workshop)
Module 7: techniques and strategies for manipulating the mouse genome (lecture only)
For each module, at least one in-depth lecture is provided. Information in the lectures is supplemented by texts and protocol manuals that are available in the laboratory where the students are working (see list at the end of this article). After the lectures for modules 1–5, detailed protocols are carried out by the students. At the conclusion of each of these modules, there is a discussion on troubleshooting. For the hands-on component of module 6, students practice using some of the databases that are available on the internet to obtain information from sequence data. Module 7 is based solely on a lecture that is purposely scheduled at the end of the course, as this lecture is designed to tie together much of the information learned from the other modules. In addition to the modules listed, two additional lectures are provided: one to outline the use of cDNA library screening and the other to discuss the various methods for analyzing differential gene expression. Here, we provide a brief outline of the lectures and a description of each module. Given that one of the modules requires the use of radioactivity, a radioisotope handling workshop is provided prior to that module. This workshop is conducted by the University’s Department of Occupational Health and Safety. In addition to providing the necessary safety training for the use of radioisotopes in the course, all of the students become officially certified to use radioisotopes in the future. This is especially beneficial because most students enrolled in the course are incoming graduate students, some of whom would be required to take the radiation safety workshop prior to beginning their own experiments. The following is a brief description of each module.
Module 1: DNA: properties and common molecular biological terminology; cDNA subcloning. The lecture component of module 1 describes recombinant DNA cloning and its uses. Techniques such as restriction digests, alkaline phosphatase treatment, ligation, and transformation are explained in detail. The lecture also discusses the components of plasmids and considerations for choosing a suitable vector. In the laboratory, students use the cloning procedures outlined in the lecture to move a cDNA from one vector to another. The students transform their new plasmid DNA into competent Eschericia coli (E. coli) cells, and, using a previously prepared transformation, they pick colonies to inoculate Luria broth for subsequent small-scale plasmid preparations. After the plasmid preparations, the orientation of the gene within the new vector is determined by restriction analysis. The importance of controls for ligations is discussed along with methods of optimizing transformation efficiency and maximizing DNA yields from small-scale preparations.
Module 2: gene transfer and protein expression analysis. Module 2 begins with a lecture describing the various methods of gene transfer into eukaryotic cells and the differences between transient and stable transfections. In addition, the various methods of analyzing protein expression are outlined, with an emphasis on Western blotting. In the lab, students transiently transfect both a control plasmid and a plasmid containing a cDNA of interest into COS P7 cells. The cDNA is fused to another cDNA encoding enhanced green fluorescent protein (EGFP), so that students can view cells under a fluorescence microscope to determine their transfection efficiency. To analyze expression further, cell lysates are collected and used to prepare a Western blot by use of an antibody recognizing the protein of interest. Postlab discussions focus on the critical steps in Western blotting for maximizing the signal-to-noise ratio. In those instances where no signal is detected, the students are taken through the various checkpoints for following the efficiency of their protocols. For instance, green-fluorescing cells were observed under the microscope, indicating that the transfection was effective, protein gels were stained with Coomassie after transfer to confirm that protein was run into the gel, and the protein marker was detected on the nitrocellulose membrane to demonstrate successful transfer.
Module 3: RNA expression analysis. To introduce module 3, the properties of RNA are introduced, along with a brief comparison among mRNA, tRNA and rRNA. After an in-depth comparison among the different methods of mRNA detection (ribonuclease protection assays, Northern blotting, RT-PCR, and in situ hybridization), the principles of RNA isolation are outlined. In the lab, students isolate total RNA (our tissues of choice have been mouse organs and Xenopus embryos) and then use both spectrophotometry and gel electrophoresis to analyze the quality of that RNA. Some of the isolated RNA is used to carry out Northern blot analysis, whereas the remainder is used in module 4. The various precautions to avoid RNA degradation are outlined, and the importance of efficient transfer to membranes and sufficient blocking and washing of the blot is discussed.
Module 4: RT-PCR. An extensive background is provided to explain the principles behind reverse transcription and the polymerase chain reaction. Students are given an appreciation for the power of this technique as a very sensitive approach to studying specific mRNAs. In addition, some time is spent discussing the various components of successful PCR including appropriate primer design and tests for authenticity of the products. Students use the RNA isolated in module 3 for reverse transcription to generate cDNA followed by PCR to obtain amplicons. To confirm the authenticity of the PCR product, a restriction endonuclease digest is carried out. Throughout this procedure, several controls are used, and sterile technique is reinforced to impress upon students the sensitivity of RT-PCR, which can easily lead to false positives. After the RT-PCR protocol, another lecture is presented to outline specific uses of this technique, including semiquantitative and quantitative RT-PCR.
Module 5: in situ hybridization. After an introduction to the principles and uses of in situ hybridization (ISH), the various steps that are critical in ISH protocols, such as permeabilizing the tissue, hybridizing probe, etc., are outlined. The method of in vitro transcription to generate riboprobes is discussed, and isotopic (32P- or 35S-labeled) and nonisotopic (digoxygenin) riboprobes are compared. In addition, the various preparations such as whole mount embryos or organs, sections, and cultured cells that can be used for ISH are discussed. Information derived from ISH is compared with that acquired from other methods of analyzing mRNA expression. In the lab, students generate digoxygenin-labeled sense and antisense riboprobes to detect the expression of mRNA encoding collagen type II (col II), an abundant marker of cartilage formation, in whole embryos and in sections of embryos. The use of both preparations teaches students the different kinds of information each can provide. For instance, the whole embryo in situs illuminate general regions of expression, such as the developing skeleton in the case of col II. In this respect, whole mount preparations are useful when expression data for a particular gene are relatively limited. On the other hand, to study the specific cell types of the developing skeleton that express col II, it is necessary to use sections. Throughout the ISH procedures, most of the discussion centers around the various methods of eliminating background signal while enhancing detection of the mRNA of interest.
Module 6: sequencing and use of sequence databases. To begin this lecture, the reasons for sequencing DNA are discussed, and the Maxam and Gilbert method of sequencing is described and compared with the more recent Sanger chain termination method. The automation of the Sanger method by the use of fluorescent-tagged ddNTPs, a laser, and computer technology is also outlined. After a description of the Human Genome Project, the two main sequencing databases, the NCBI database (www.ncbi.nlm.nih.gov/) and the TIGR database (www.tigr.org/tdb/tdg/html), are described. After the lecture, students are given sequence autoradiograms to decipher, and they use the databases to identify their genes. The sequences given to the students are fragments of genes whose full-length clones have not yet been characterized. Thus sequences can be blasted in GenBank, and homologous regions can be seen; but to gain further information, students must use the TIGR database. In addition, students are taught how to use SwissProt to look at potential protein domains of their unknown sequences.
Module 7: techniques and strategies for manipulating the mouse genome. The objective of module 7, which is a lecture only, is to provide an introduction to the techniques currently available for manipulating the mouse genome. The generation of transgenic mice is discussed first, followed by directed mutagenesis in embryonic stem (ES) cells. The applications of transgenic technology are outlined, including investigating the basic mechanisms of tissue-specific or developmental stage-specific gene regulation, generating mouse models to study human disease, and studying pathophysiological conditions. In addition to describing construct design and microinjection into the pronucleus of fertilized oocytes, a major technical concept introduced is the use of tissue-specific promoters to provide spatial and/or temporal control over transgene expression. Students are also introduced to the use of "promoter bashing" to identify minimal promoter regions.
To introduce gene targeting strategies, a background on ES cells, chimeras, blastocyst injection, and aggregation is provided. The technical background provided includes the generation of basic targeting constructs and methods of screening for homologous recombinants. This is followed by a discussion on the strategies for greater control of loss-of-function and gain-of-function mutations, including the generation of conditional knockouts and the use of inducible systems to control transgene expression. Although very in-depth, this lecture is very effective in fulfilling two major objectives. First, the lecture serves to allow students to put all of the techniques they have learned into a greater context as they become aware of how each of these techniques is critical for generating and analyzing transgenic or knockout mice. Second, students are introduced to state-of-the-art techniques for analyzing gene function, techniques they will frequently encounter in research articles and seminars.
ADDITIONAL LECTURES
Library screening. A lecture is provided that introduces students to cDNA libraries and their uses. Procedures for making cDNA libraries are outlined, and the differences between random-primed and oligo (dT)-primed libraries are discussed. The technique for screening cDNA libraries is also explained, and students are shown sample autoradiograms from a library screen.
Approaches for identifying developmentally or physiologically important genes. This lecture is a recent addition to the course, and arose from our realization that, in the past, students were unaware of situations in which one would obtain a small, unknown piece of DNA. To address this, a lecture was designed to outline the different methods of examining differential gene expression. The methods are covered in detail with respect to how they work and the advantages and disadvantages of each technique. The common methods that are discussed are the candidate approach, differential display analysis, cDNA expression arrays, subtractive hybridization, and microchip arrays. Throughout this lecture, emphasis is placed on the transition from "hypothesis-based" research to "discovery-based" research that some of these techniques involve.
COURSE SCHEDULE
After offering this course for four consecutive years, we have established a schedule (Fig. 1) that is effective at meeting the time requirements for each module while limiting the "down-time" that is often associated with molecular biology protocols. The following is the schedule for days 1 and 2 of the course as an example of how the modules are interleaved to make efficient use of time.
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08:30 - introduction to the course; expectations
09:00 - begin module 1 with lecture on DNA: properties and common molecular biological terminology; restriction endonucleases and ligases; introduction to restriction mapping and subcloning; considerations for choosing a vector
10:00 - set up restriction endonuclease digests for subcloning; prepare agarose gel for separating fragments
11:00 - lecture on library screening; add alkaline phosphatase just before lunch
12:00 - lunch
13:00 - load and run gel; prepare ampicillin plates; gel purification of insert; set up ligation and overnight cultures for mini-preps.
15:00 - begin module 2: transfection of COS cells with expression construct
17:00 - cleanup and questions
Day 2:
08:30 - isolate mini-prep DNA and set up digestions to determine orientation of inserts; pour gel and start restriction endonuclease digests
12:00 - lunch
13:00 - run gel, during which time transformations with students’ own DNAs will be set up; interpret results from gel while transformed cells are incubating
15:30 - plate transformations
16:00 - lecture on gene expression and analysis of protein expression
17:00 - view transfected COS cells with fluorescence microscope
17:30 - cleanup and questions
SPECIAL CONSIDERATIONS
The smooth running of this course has always relied on having an assistant coordinator to oversee the organization and protocols and to run the labs. In the Department of Physiology, this role is filled by a senior graduate student with extensive experience and background in all of the techniques covered in the course. This individual is typically responsible for making the required solutions (with the help of a departmental technician) and for working with the faculty members to optimize the protocols and acquire all of the required materials. The assistant coordinator is present, along with the faculty organizer, for each module during all of the lab procedures to answer questions, demonstrate techniques, and help students with troubleshooting.
Another consideration we had while organizing the practical aspect of the course was which genes to use for each protocol. In the first year of offering the course, the same cDNA was used throughout all those modules for which a cDNA was required (cloning, protein expression analysis, Northern analysis, RT-PCR, and in situ hybridization). Originally, it was thought that using the same cDNA throughout the entire course would provide continuity and avoid confusion. However, we later opted for having each faculty member choose the cDNA for her/his own modules. By working with cDNAs that they are familiar with and that they have had success with in their own research, faculty members are better able to design protocols that run smoothly.
A major concern in establishing a course such as this is the expense of the reagents and equipment that are required to provide students with the hands-on experience that makes this course so valuable. In building this course from the ground up over that past four years, this has been one of our greatest challenges. An initial commitment from the Department of Physiology was required to provide funds for buying the reagents and equipment that were absolutely essential for offering the course. In addition, laboratory space adjacent to an already established core molecular biology facility was provided for the three-week duration of the course. Among the facilities available in this area are thermal cyclers, a fume hood, a refrigerator, a freezer, a hybridization oven, centrifuges, incubators, and a separate room designated for the use of radioactivity. Other major pieces of equipment already owned by the department include a gel imaging system, a transilluminator, and computers, all of which are within close proximity and are made available for teaching purposes. In this respect, cooperation from all members of the department has been essential. Initial investments included the purchase of several reagents, power supplies, gel apparatus, hot plates, and vortexes. Over the past three years, we have been able to purchase additional equipment and eliminate the need for sharing things like gel boxes among more than two students. In comparing the fourth and most recent version of the course to the first offering, we have found that, although it may seem inefficient economically, the less sharing done between groups of students the better. This enables pairs of students to work at their own pace while enhancing the hands-on experience. Nonetheless, offering the course to eight students requires no more than two power supplies, two vortexes, one centrifuge etc.
The cost per student for expendable supplies is estimated to be $500. A major portion of this comes from the use of enzymes and reagents for reactions such as labeling riboprobes and carrying out RT-PCR and restriction endonuclease digestions. Whenever possible, we take advantage of surplus reagents from the labs of participating faculty. However, the successful outcome of most protocols relies heavily on obtaining fresh enzymes; thus this expense is essentially unavoidable. To offset some of these expenses, supervisors of course participants who are not graduate students in our department (i.e., graduate students from other departments or technicians/postdocs) are charged a $500 fee. Despite our best efforts at economizing, this course is expensive to offer; however, given the very positive outcome of the course, we feel that this expense is justified.
STUDENT EVALUATION
The primary objective of the course is to provide students with an understanding of how they can use molecular techniques to address a variety of questions and to provide them with the basic skills to carry out those techniques. To evaluate students’ comprehension of course material, a written final exam is prepared in which each faculty member provides questions on his/her module. Some exam questions require the students to describe how they would use the techniques learned to address a particular problem, whereas others may ask students to compare two methods and describe advantages and disadvantages to each. The format of the questions is left to the discretion of each faculty member. As an example, the following question was designed to test the students’ understanding of the RT-PCR module:
You are studying a peptide hormone in sheep that you know is synthesized in the pituitary gland. The cDNA corresponding to that same hormone has been cloned from rat, mouse, and human tissues. Using the rat cDNA, you probe a Northern blot of poly(A)+ RNA from the whole sheep pituitary gland and obtain a single, faint band on the autoradiogram. How could you use RT-PCR to confirm that this is the mRNA you were looking for? How would you identify the specific cells producing the hormone?
The final exam appears to provide an effective measure of the students’ understanding of the course material; however, relying solely on this exam for evaluation has met with criticism from students, some of whom feel the exam does not appropriately reflect their efforts in the course. In addition, we have come to realize the importance of evaluating student progress continually during the course and therefore have begun to add quizzes to the modules. The quiz questions may test specific technical knowledge or pose troubleshooting problems. The aim is to help students stay abreast of the material as the course proceeds. The quizzes have the added benefits of serving as a measure of how students are progressing and helping to identify when and where students require extra help.
COURSE EVALUATION
All participants in the course have been extremely satisfied with the outcome. The general consensus is that this is a unique and beneficial opportunity. One of the aspects of the course that students comment on from year to year has been the efficient course organization: the intensive, three-week format before the start of the autumn term allows them to minimize scheduling conflicts with other courses. In addition, students enjoy the interaction with various faculty members and recognize how fortunate they are to have the course offered within their own department. Although some of the techniques taught in this course may have been introduced in one or another of their undergraduate courses or through summer jobs, having the material presented as an integrated "package" encourages a better appreciation for the relative strengths and weaknesses of the various techniques. On the other hand, for those with no prior experience with molecular techniques, this course provides a thorough introduction that integrates theory and practice. Indeed, we have received positive feedback from faculty members whose students completed the course. One of our overall goals is to equip students working on their individual projects with the skills and expertise to repeat the protocols in their own labs, and this course has been successful for four years in meeting this goal. Importantly, it does so early in the student’s graduate program and at a cost to the supervisor that is much less than the cost of sending a student to another institution for training.
One of the main reasons for the success of this course has been that, upon its completion each year, the course coordinator meets with the students as a group to receive their comments. This session has been an important source of ideas for improving the course in succeeding years. In addition, starting in 2001, the course and its participating faculty are being formally evaluated by the University’s system of computer-automated course and instructor evaluations. The results of that first formal evaluation were not yet available at the time this article was prepared.
Acknowledgments
Address for reprint requests and other correspondence: G. M. Kidder, Dept. of Physiology, Univ. of Western Ontario, London, ON, Canada N6A 5C1 (E-mail: gerald.kidder{at}fmd.uwo.ca).
Received for publication June 24, 2001. Accepted for publication December 5, 2001.
REFERENCES
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