Intensive Laboratory in Biotechnology 
Carol Lin

 

May 19 - July 10, 2014, plus one take home exam due a week later.

M - R, 9 am - 2 pm, plus additional independent lab work as necessary.

744E and 743 Mudd

*** First day of class is May 19, 9 am, in 744E Mudd. Come even if you have not registered.***

 

Rationale

Hands-on exercise is one of the most important aspects in experimental biology.  Students enrolled in Biotechnology programs have an extensive opportunity to learn many cutting-edge molecular biology methods from lecture- and reading- based courses.  Laboratory lectures.  Students will learn practical skills in planning, executing, analyzing and troubleshooting research protocols.  The goal of this intensive laboratory is to expose students to various techniques in biotechnology as well as to prepare them for independence in research settings.  To ensure the continuation of experimental protocols and simulation of an actual research situation, classes are scheduled to meet long and frequently.  With such an immersion in the laboratory, students will learn not only a comprehensive array of techniques, but also the real-life issues that are frequently missing from shorter laboratory courses.  This course is designed for student who do not have any prior independent research experience. 

 

Course content

Students working in teams of 2 – 4 students will meet four 5-hour days a week for eight weeks during the summer.  The course will concentrate on modern laboratory methods in biotechnology.  These include the engineering and analyses of genes and gene products (RNA and proteins) in bacteria, yeasts, and mammalian cell culture. 

 

Most class meetings will begin with a recitation period when background information, experimental design, calculation, and task scheduling will be discussed.  Actual laboratory will include all steps in conducting experiments.  These may include reagent preparation, equipment set-up and calibration, protocol running and monitoring, laboratory maintenance, accident prevention, simulated accident handling, post-lab clean up and waste disposal.  Each laboratory exercise will conclude with data analysis, discussion, and presentation.

 

Students will work in small groups and will be expected to forge collaboration within and among groups. 

 

Reading list

There is no text book.  Reading materials will be distributed during class.

 

Course requirements

 

Syllabus

Number of days for each topic is shown in parentheses. 

Different topics may be held concurrently.

  1. Introduction, general procedure, and lab safety
  2. Manipulation DNA

    a) Large scale plasmid DNA preparation

    b) Restriction enzyme digestion and gel electrophoresis

  3. Subcloning

    a) Gel purification

    b) Ligation

    c) Preparing competent cells

    d) Transformation

  4. Screening

    a) Plasmid mini preparation and gel analysis

    b) Probe generation

    c) Southern transfer

    d) Southern hybridization

    e) Detection

    f) Colony hybridization

    g) PCR analysis

  5. Site-directed mutagenesis as Practical Exam I
  6. Tissue culture technique

    a) Maintenance, transfection and selection

    b) High molecular weight DNA isolation

    c) RNA isolation

  7. Gene expression analysis

    a) cDNA generation

    b) Array analysis

  8. Yeast two-hybrid analysis

    a) Fusion gene construction

    b) Yeast transformation

    c) Phenotype testing

  9. Protein analysis

    a) Expression and purification

    b) Western blot analysis

  10. Expression analysis in eukaryotic cells as Practical Exam II

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