Dr. Craig Pikaard: pikaard@biology2.wustl.edu

Course syllabus (PDF)

Overview of the course:

This course, offered in partnership between the Washington University Biology Department and the Donald Danforth Plant Science Center (DDPSC), is designed for beginning graduate students and advanced undergraduate students contemplating a research career. The course focuses on methods for the biochemical analysis and imaging of plant proteins. Topics include measurement of protein concentrations, purification of recombinant proteins, assessment of protein purity by SDS polyacrylamide gel electrophoresis, protein identification using mass spectroscopy, protein crystallization, and an introduction to protein structural analysis. Students also transform plant cells in tissue culture in order to express recombinant fluorescent proteins that are later visualized within living cells using confocal fluorescence microscopy. The course meets for one hour on Monday afternoon, for a lecture covering the concepts for the week's laboratory sessions, and for two 4-hour laboratory sessions on Wednesday and Friday afternoons, respectively.

Course Description

This course focuses on methods for the genetic engineering, manipulation and analysis of proteins. We begin by purifying a thermostable DNA polymerase overexpressed in E. coli, taking advantage of the thermal stability of the protein as well as ion-exchange chromatography in our purification scheme. Purity is assessed by subjecting fractions from each purification step to SDS polyacrylamide gel electrophoresis and then staining the resolved proteins using the protein-binding dye Coomassie Blue. In parallel, we test for activity of the polymerase using a polymerase chain reaction (PCR) assay. Combining the two assays allows students to identify a specific protein band that corresponds to the detection polymerase activity.

The students then use the polymerase they have purified to clone, by the process of reverse transcription-polymerase chain reaction (RT-PCR), the coding regions of genes encoding proteins of current research interest. The cDNA clones are captured in Gateway entry vectors and resulting clones are verified by restriction endonuclease mapping and DNA sequencing. The cDNAs are then recombined into two distinct destination vectors designed for expression of the proteins in either bacteria or plants. The genes cloned into the bacterial vector are used to overexpress the proteins in E. coli. The overexpressed proteins are gel-purified, verified by mass spectrometry and then sent off for the production of antibodies. Although the resulting antibodies are not available until after the end of the semester, the students have the opportunity to do experiments involving other antibodies, including immunoprecipitation and immunoblot (western blot) detection of epitope-tagged proteins.

The cDNAs cloned into the plant destination vector generate fusions of the encoded proteins to yellow fluorescent protein (YFP). These vectors are transformed into Agrobacterium and resulting cultures are infiltrated into tobacco leaves. 48-72 hours later, the students use fluorescence microscopy to examine the subcellular localization patterns for their proteins.

In a module taught by Dr. Joe Jez, students set-up a matrix of buffer and salt conditions for the crystallization of lysozyme using the hanging-drop approach. Students record the progress of the crystallization process and ultimately select crystals, mount them in the X-ray beam and obtain X-ray diffraction data and images. Although there is not time to have the students solve the structures, the students are taught the steps involved in the process and learn to use PyMol software in order to examine existing crystal structures and answer structure-based questions that comprise a problem set.

Laboratory notebooks- advice and expectations for students
Buy a bound (not a loose-leaf or 3-ring binder) notebook with numbered pages to serve as your graded laboratory notebook. You will also want a separate notebook for storing lecture notes and handouts; a 3-ring binder will probably be useful in this capacity.

1) Label the first page of the Laboratory notebook: Table of contents.
On this page you will list the various experiments and the page numbers where the relevant experimental procedures and data can be found. This will help you and the instructors locate information quickly.

2) Use a ball-point pen for all notes, not pencil (or ink that will run if wet).

3) Each experiment begins on a new page (on the left side of an open notebook) and gets an experiment number at the top of the page. The experiment number is composed of your initials and the dates, e.g CP070119 would be the experiment number for Craig Pikaard on Jan 19, 2007.
-Any tubes, gels, data etc. pertaining to a given experiment will be labeled with the appropriate experiment number, even if the data are gathered on a subsequent day.

-use the left-side page for your working notes while the lab is in progress, keeping each right-side page blank. Later, you will copy (neatly) and edit (if necessary) your notes onto the right hand side page. Grading will be based on what is written on the right side.

4) Following the experiment number, the next item on the page is the experiment name - ask your instructor for what this name should be.

5) Next comes Objectives: here you state the purpose of the experiment in a sentence or two.

6) Next comes References: refer to any protocols, publications (or previous experiment numbers elsewhere in the notebook) that pertain to the experiment

7) Next comes Procedures:  here, you provide your step by step descriptions of the experiment(s). Most likely, this section will also include results as you obtain them and move forward with subsequent steps.

8) Finally comes the Summary of results and Conclusions: did the experiment work? If so, what were the findings and what do they mean (interpretation). If not, what do you think might have gone wrong and what corrections should be tried if the experiment were repeated etc.