BNAT home
BNAT course
Fluorophores and Microscopes
The faculty of the Basic Neuroscience Advanced Training (BNAT) program offer a course called “Fluorophores and
Microscopes.” The course features classroom sessions (lectures, student and
faculty presentations) and didactic and independent laboratory work (utilizing
the four advanced microscopes in the Light Microscopy Facility). This
course is designed to provide both a theoretical background and practical
experience for students in fluorescence microscopy and the use of exogenous and
genetically engineered fluorophores.
Who: BNAT
trainees and other PhD students, postdoctoral fellows, and faculty. Enrollment
limited to 12.
When: Tue
& Thu, 1.5 hrs each meeting; Spring semester,
2006. 29 sessions total. Time of day TBA.
Where: Neuroscience /Physiology & Biophysics
conference room, RC1 north, 7th floor
Text:
There is no required text. Copies of relevant textbooks will be made available
as appropriate. Handouts, online materials, and original papers will also be provided.
Evaluation
of students (by faculty) is according to performance on individual laboratory
projects and classroom and laboratory participation. There are no written
examinations. Evaluation of faculty (by students) is according to clarity of
presentations, appropriateness of content, availability for and helpfulness of
consultation.
Faculty:
BNAT trainers: Drs. Beam, Betz, Ribera, Caldwell, Dell’Acqua,Finger, Levinson,
Restrepo, Sather; BNAT trainee Dr. Joe Johnson, non-BNAT faculty: Dr. Nicholas Barry, Assistant Professor, Dept. of Medicine; Dr. Amy Palmer, Assistant Professor, Chemisty & Biochemistry (Boulder campus) Teaching
Assistants: BNAT trainee Dan Sdrulla;
Former BNAT trainee Dr. Ernie Salcedo; Light Microscopy Facility Manager Fadul;
Other trainees Dr. Sophie Breusegem, Michael Gaffield, Marc Yonkers.
Content:
The course is divided into seven parts. Part 1 is an overview of the entire
course. It is largely didactic and relatively highly structured. Part 2 gives
hands-on experience with advanced microscopes in the Light Microscopy Facility
(LMF) or Electron Microscopy Facility (EMF). Part 3 is back in the classroom for discussion of special techniques. Part 4 involves faculty research
seminars (presenters chosen by students). The focus is on advanced techniques (faculty lectures and student presentations). Parts 5-7 involve advanced topics and student lab projects that focus on technology, not biology, although students may use preparations of their choosing.
Administration. Mrs. Andrea Banks, UCD at Fitzsimons, RC1 North Tower Room P18-7130. voice: 724-4500. email: andrea.banks@uchsc.edu
|
Schedule: FLUOROPHORES AND MICROSCOPES
|
|
PART I: Course
overview. The first session,
students will tour the LMF, which is directly adjacent to the classroom. The
next 4 sessions will be an overview of the course, and provide the basic
tools for understanding the sessions that follow.
|
|
Jan 24
|
Betz/Fadul
|
Introduction, tour of LMF (microscope demos
by senior trainees)
|
|
Jan 26
|
Beam/Betz
|
Optics (diffraction limit, resolution,
illumination, image formation, filter sets), Microscopes (conventional, laser scanning confocal,
digital deconvolution,
two photon, TIRF, STED-4pi)
|
|
Feb 2
|
Levinson/Johnson
|
Fluorescence (chemistry
of fluorophores, excited state, emission and excitation spectra,
photobleaching, other fluorescence phenomena)
|
|
Feb 7
|
Levinson/Palmer
|
Fluorophores (exogenous dyes, immunofluorescence,
genetically encoded fluorescent proteins)
|
|
Feb 9
|
Restrepo/Barry
|
Image acquisition (CCD cameras, laser scanners), Image processing (enhancement, feature extraction)
|
|
PART II: Laboratory
hands-on experience. Students
will divide into 4 groups and rotate each week for 4 weeks to the 4
microscopes in the LMF. The objectives are to become familiar with using each
instrument, and to understand in detail the principles by which they operate.
A teaching assistant or faculty member will assist at each microscope. See www.uchsc.edu/lightmicroscopy
for a detailed description of the microscopes.
|
|
Feb 14
|
Lab
|
The 4 microscopes
are:
•
Zeiss 510 NLO
(two photon) Meta; HP Fluorimeter
•
Olympus
TIRF
•
Deltavision
Digital Deconvolution
•
Olympus
spinning disk
|
|
Feb 16
|
Lab
|
|
Feb 21
|
Lab
|
|
Feb 23
|
Lab
|
|
PART III:
Techniques. In these classroom
sessions, the major techniques by which fluorophores are created and used
will be discussed and (in some cases) demonstrated.
|
|
Feb 28
|
Caldwell/Sather
|
FRAP, FLIP,
exogenous fluorophores
|
|
Mar 2
|
Sather/Beam/Dell’Acqua
|
FRET
|
|
Mar 7
|
Barry/Beam
|
FCS, FLIM (demo and
discussion at 9th Avenue Campus (Moshe Levi’s lab – 4th
floor, BRB)
|
|
Mar 9
|
Dell’Acqua/Ribera
|
Fluorescent
proteins, tour of transgenic core facility
|
|
Mar 14
|
TBA
|
Other (e.g., 2nd
harmonic generation, calcium imaging, quantum dots)
|
|
PART IV: Faculty
research presentations. Faculty
will present research seminars that illustrate the use of these optical techniques.
Faculty will be selected by the students.
|
|
Mar 16
|
TBA
|
|
|
Mar 28
|
TBA
|
|
|
Mar 30
|
TBA
|
|
|
Apr 4
|
TBA
|
|
|
PART V: Student
presentations. Each day for 4
sessions, 3 students will present an advanced topic, a journal article, or a
proposal for his/her independent laboratory project. Each will rehearse with
a faculty member, who will be present during the presentation.
|
|
Apr 6
|
Student
presentations
|
|
|
Apr 11
|
Student
presentations
|
|
|
Apr 13
|
Student
presentations
|
|
|
Apr 18
|
Student
presentations
|
|
|
PART VI: Student
lab projects. Students will work
on projects, either independently or in teams, using equipment in the LMF (or
electron microscopy facility). Projects can be independently proposed, or
chosen from a list. NOTE: Work does not have to be confined to class
schedule, since the LMF is open 24/7, and students will be qualified users of
the instruments.
|
|
Apr 20
|
Lab projects
|
Potential Projects
(students may also design independent projects)
•
Point-spread
function: Measure the PSF of two
microscopes in the LMF
•
Bleedthrough: Given fixed material that is dually
stained, measure the amount of emission ‘bleedthrough.’ Compare results with
that obtained by using the Zeiss Meta.
•
FRET: Measure FRET of cameleon transfected
into tissue culture cells.
•
FCS: Measure the diffusion coefficient of GFP
in tissue culture cells (9th Ave campus), or fluorescent beads
in media of different viscosities.
•
TIRF: Compare the Brownian movements of
fluorescent beads of different sizes. Measure the airy Airy disk of
diffraction-limited beads.
•
Confocality: Measure the effects on xy axis and z axis
resolution by adding or removing the nipkow Nipkow disk
in the Olympus spinning disk microscope.
•
Image
processing: Given an image stack,
examine methods of feature extraction (e.g., nodes of Ranvier, secretory
granules).
•
Photobleach: Compare the effects of various anti-fade’
agents (e.g., Profade, Vectashield,Prolong on
bleaching of fluorophores
•
Meta (Zeiss) versus fluorimeter: Measure
emission spectra with both instruments and compare results
•
2-photon imaging: image GFP transfected motor neurons in zebrafish embryos
|
|
Apr 25
|
Lab projects
|
|
Apr 27
|
Lab projects
|
|
May 2
|
Lab projects
|
|
PART VII: Student
presentations. Students will
present and discuss results from their laboratory projects (4 presentations
per session)
|
|
May 4
|
Student
presentations
|
Students will
analyze results from their laboratory projects, rehearse with appropriate
BNAT faculty, and then present their work in class.
|
|
May 9
|
Student
presentations
|
|
May 11
|
Student
presentations
|
|