Principal Investigator/Program Director Williams,Robert W.
A (Neurohistology Core)
The specific objective of the Neurohistology Core is to provide high-quality histology for several aspects of the research comprising the Program Project. Specifically, this work includes the basic histologic processing of brains to be part of the Mouse Brain Library (MBL, Project 1), which will be used by Project 2 (iScope) and Project 3 (Neurocartographer).
The Neurohistology Core will be responsible for ensuring proper collection, registration, and processing of all neuroanatomic materials generated by the component projects.
In consultation with other investigators of the Neuroinformatics Program Project, the Neurohistology Core will be involved in planning, application for funding, and implementing additional research utilizing neuroanatomic and pathologic analysis that can be attached to the current and future research activities of the Program Project.
The Neurohistology Core will be staffed by one full-time professional and one full-time and one half-time histology technician.
attached biographical sketch)
Dr. Glenn D. Rosen, Ph.D., Principal Investigator of the Neurohistology Core (10% effort), will supervise all activities of the Core.
B. Support Personnel
Stefany Palmieri, Head Technician, will histologically process all brains for the Neurohistology Core. Ms. Palmieri has been involved in processing the 600 brains currently in the MBL. She is highly skilled in all aspects of processing brains for celloidin embedding. In addition, we will hire a technician who will split his/her time between the Neurohistology Core and Project 1. This person will aid Ms. Palmieri in processing the brains and will be responsible for imaging these brains for uploading to the MBL.
3. Resources and Environment
The Neurohistology Core will
be housed at the Beth Israel Deaconess Medical Center. This laboratory is
located on the seventh floor of the Charles A. Dana Research Building of
The laboratory contains approximately 2100 square feet, 1500 of which are wetlab space. Half of this space will be devoted to Neurohistology Core activities. The space includes a fully equipped histology area housing a giant microtome capable of processing wholebrain human specimens, sliding microtomes, a rotary microtome, a cryostat, an oven, a refrigerator, a rocker table, a staining microscope and accessories, scales, meters, and glassware. There is a microscopy room containing a teaching comparison compound microscope, two stereomicroscopes, a compound light microscope, and equipment for macro- and microphotography. Additional facilities include cabinets for storage of serial histologic sections and a fully equipped darkroom for photographic and autoradiographic processing.
We have requested and have received commitments from Beth Israel Deaconess Medical Center for an additional 500 sq. ft. of research space allocated to us (see attached letter).
The technicians will report to Dr. Rosen. The staff of the Neurohistology Core will participate fully in the administrative and research meetings organized by the Administrative Core and specified elsewhere on this application. In addition, the Core Director will review materials for quality on a daily basis.
Dr. Rosen will be notified personally during a research meeting, by telephone, or by email that specimens will be shipped at a particular time and date. Dr. Rosen will alert the technical staff in the laboratory and the receiving department of the hospital. In some cases, specimens will be obtained directly by a member of the lab at the site of origin. The specimens will be received by the head technician and each brain will be assigned a unique identifying number.
Funds for the Neurohistology Core will be managed by the Office of Research Administration of the Beth Israel Deaconess Medical Center as part of theNeuroinformatics Program Project management. Additional organizations involved in the grant administration include the Department of Neurology, Beth Israel Deaconess Medical Center.
Interaction among the Administrative Core and other Program Project cores, the different research components, and the Neurohistology Core will be facilitated by an electronic mail network as well as facsimile facilities.
The Neurohistology Core is necessary because a large number cases will be added to the MBL. Because tissue processing is repetitive and is not in itself hypothesis driven, it is best carried out in a core facility. A centralized Neurohistology Core will facilitate the acquisition of material for the MBL, thus ensuring that the potential of the Neuroinformatics Program Project research can be realized. The Core will process and analyze neuroanatomic materials from all component projects, obviating the need to duplicate expensive equipment and train more staff in time-consuming, difficult techniques (described below).
The Neurohistology Core is
equipped to process material of varying size, from the human brain to the
brain of a mouse embryo. Personnel in this facility have carried out
studies on the brains of apes, dolphins, old world monkeys, rats, and
mice, and members of the staff have experience with a large number of
modern neuroanatomic methods applicable to the mammalian brain. See below
for details on a sampling of available techniques and protocols.
A. Routine Histologic Procedures
In preparation for routine examination, animals are perfused transcardially by the Genotyping and Mouse Colony Core (Core B), then shipped to the Neurohistology Core. Once the brains arrive in the laboratory, they are given a unique identifying number and are randomly assigned to be cut in either the horizontal or coronal plane. After post-fixation for at least one week in 10% formalin, the brains are dehydrated in a series of 80%, 95%, 100% ethanol and ethanol/ether. The brains are then placed into 3% celloidin for one week followed by 12% celloidin for 23 days or until hard.
The celloidin block is trimmed to achieve a stable base and is notched on the left side for side orientation. The specimen is placed on a sliding microtome. The sections are cut coronally at 30 m and are segregated and saved in 80% ethanol. Every fifth section is stained for Nissl substance. Spare sections are preserved indefinitely in 80% ethanol.
For the 3D atlas described in Project 3, we will cut the tissue at 10 m.
Sections are washed in distilled water and placed in 1% cresyl violet acetate solution (which stains the Nissl substance) for 35 min. Each section is placed in distilled water for 1 minute and then differentiated and dehydrated in 70, 80, and 95% ethanol. A few drops of colophonium are added to the 95% ethanol baths. If differentiation is adequate, the sections are then cleared with terpineol and passed through xylenes. Sections are mounted with careful attention to orientation so that left and right are consistently identifiable. Coronal sections are mounted with the nick in the celloidin on the right side of the slide. Horizontal sections are mounted with the nick on the left side of the slide. The sections are then mounted in Permount. One series of every fifth section will be sent to Project 1 (MBL) and subsequently to Project 2.
The MBL currently is comprised solely of Nissl-stained sections. The laboratory is prepared, however, to stain adjacent sections of celloidin-embedded material using the following methods.
Myelin Staining (Loyez). Free-floating sections are washed in distilled water for 30 s and placed for 6 h in a 2% ferric ammonium sulfate solution. The sections are then washed for 30 s and incubated overnight at room temperature in a 1% hematoxylin solution in 10% ethanol/2% lithium carbonate. The next day the sections are washed twice for 30 s each and differentiated in 2% ferric ammonium sulfate until the gray matter appears. The sections are then washed in three changes of distilled water for 30 s each before differentiation in Weigerts solution (2% sodium borate/2.5% potassium ferricyanide). The sections are then washed three times in distilled water, with the second wash containing a few drops of ammonium hydroxide, before being mounted onto subbed slides, dehydrated, cleared with xylenes, and coverslipped with Permount.
Hematoxylin and Eosin. This routine neuropathological stain identifies nuclei, cytoplasm, and blood vessels. Free-floating slides stored in 80% ethanol are placed in hematoxylin for 5 min, then differentiated in 1% acid alcohol. Each step is followed by a wash in distilled water. Then the slides are placed in distilled water containing a few drops of ammonium hydroxide (bluing agent) followed by 1 min in eosin. Then the slides pass through a series of 70%, 80%, and 95% ethanol and are stored in alpha terpineol until coverslipped.
PTAH. This routine neuropathological reagent stains astrocyte gliosis. Each step in this protocol is followed by a wash in distilled water, except where noted. Free-floating slides stored in 80% ethanol are incubated in saturated mercuric chloride for 3 h and then placed in Lugols iodine for 5 min. The sections are placed in 95% ethanol, followed by 0.25% potassium permanganate for 5 min and 5% oxalic acid for 5 min. The sections are washed 5 times in distilled water and placed in PTAH for 24 h. The sections are washed in 100% ethanol, placed in xylenes, and coverslipped.
Massons Trichrome. This stain identifies
nuclei, collagen, and blood vessels. Each step in this protocol is
followed by a wash in distilled water, except where noted. Free-floating
slides stored in 80% ethanol are placed in Weigerts iron hematoxylin
for 30-45 s and then in Massons fuschin OG for 5 min. Sections are then
placed in 1% acetic acid for 3 min, 5% phosphotungstic acid for 5 min, 1%
acetic acid for 3 min, 2% light green for 5 min, and 1% acetic acid for 1
min. There are no washes between these steps. Sections are mounted and
placed in 80% ethanol, 95% ethanol, and xylenes and coverslipped.
Gallyas Protocol. Because this method yields optimal contrast for image analysis, it can be used on sections from which cell counts are made. The celloidin sections are cleared of celloidin and mounted onto glass slides. The slides are then placed into 4% formic acid for 3 h and then overnight in a solution of 10% formic acid and 9% hydrogen peroxide. The next day, the slides are washed three times for 15 min each and then placed for 1520 min into the developer (2.5% sodium carbonate, 0.1% silver nitrate, 0.1% ammonium nitrate, 0.5% silicotungstenic acid, and 0.36% formalin). The sections are then placed in 0.5% acetic acid for 5 min, washed in distilled water for an additional 5 min, and fixed in Extaflo fixer (0.29% for 10 minutes). After another 5-min wash in distilled water, the sections are dehydrated, cleared, and coverslipped.
Potential Pitfalls and Problems with Celloidin Processing. A minor technical problem concerns tissue shrinkage during processing. For many CNS traits it is of interest to compute in vivo values. This can be difficult when the data are obtained from celloidin material where tissue shrinkage is severe. Our solution to this is to compute the degree of shrinkage for individual brains. Brain volume after processing is computed by using a uniform point counting protocol. This value is then compared to the brain weight obtained immediately after dissection. We have used this method in several cases and volumetric shrinkage is approximately 5060%. From these volumetric shrinkage estimates we can compute approximate linear shrinkage. We have also directly estimated shrinkage by comparing MRI estimates of brain volume with those of the same brain after processing, and have obtained comparable estimates of shrinkage.
As the MBL expands, we will begin to include immunohistochemically stained tissue. A large number of immunohistochemical stains have been implemented in the Neurohistology Core laboratory. New antibodies are continually being used in response to the needs of particular research questions. For the current proposal, we plan only to stain for ChAT‑ and Parvalbumin‑immunoreactive neurons and fibers. In the future, we may expand the MBL even further, and we therefore include protocols for some of the other candidates. Antibodies have been obtained from commercial sources and from laboratories of colleagues
For immunohistochemistry, mouse pups and adults are perfused transcardially under deep anesthesia with 0.9% saline followed by 4% paraformaldehyde. The brains are removed from their skulls and post-fixed for 24 h in 4% paraformaldehyde, then placed in a 0.1 M phosphate buffer/10% sucrose solution until the brains sink (usually 1 day). Next, the brains are placed into a 0.1M phosphate buffer/30% solution until they sink, usually 23 days later. The brains are nicked on the ventral surface of either the right or left hemisphere,then cut frozen on a sliding microtome in the coronal plane at 30 m. Consecutive sections are stored in 0.1M phosphate buffer.
Neurofilament (NF). Free‑floating sections are rinsed twice in phosphate‑buffered saline (PBS; pH 7.4) for 5 min each and transferred to a buffered 0.6% hydrogen peroxide solution in order to block staining of endogenous peroxidases. The sections are rinsed twice in PBS and incubated overnight at 4C in a 1/50 dilution of mouse anti‑neurofilament immunoglobulin (monoclonal antibody to the 68 kDa subunit of neurofilament from Boehringer Mannheim, Indianapolis MN). The vehicle (diluent) for all antibody incubations is 3% rabbit serum in PBS.
Sections are then placed into a solution containing the linking antibody (rabbit anti‑mouse immunoglobulin, Dakopatts, Santa Barbara, CA, Z259; diluted 1/20) at room temperature for 2 h. The sections are rinsed twice with PBS and placed in a 1/250 dilution of mouse peroxidase anti‑peroxidase (Dakopatts B650) at room temperature for 2 h. The tissue is rinsed twice in PBS and then twice in 50 mM Tris buffer (pH 7.6) and developed using 0.05% diaminobenzidine and 0.005% hydrogen peroxide diluted in Tris. After rinsing with Tris, sections are mounted on chrome‑alum coated slides, dehydrated, counterstained with Methyl Green/Alcian Blue, and coverslipped with Permount.
The following monoclonal antibodies are suitable for use in mouse and are also available to Core A. Note again that to maintain high-throughput we initially only intend to stain for two of the following short list of cell types (ChAT and parvalbumin).
1. Radial glial fibers (Rat-401). This monoclonal antibody directed against radial glial fibers is provided by S. Hockfield, Yale University. The protocol is the same as that for NF with the exception of the dilution (1/4) of the primary antibody.
2. Vimentin. This monoclonal antibody (Boehringer Mannheim) stains radial glial fibers with less background than Rat-401. The protocol is the same as that for NF with the exception of the dilution (1/500) of the primary antibody.
3. Glutamate. This antibody (Incstar) stains glutamatergic fibers with a few cell bodies. The procedure for glutamate staining is identical to that for NF with the exception of the dilution (1/5000) of the primary antibody.
4. Parvalbumin. This antibody (ICN) stains neuronal cell bodies in the cortex (many of which are GABA positive). The staining procedure is identical to that for NF with the exception of the dilution (1/500).
5. Choline acetyltransferase (ChAT). This antibody (Chemicon) stains fibers and cells of the ascending cholinergic system of the forebrain. The procedure is the same as that for NF with the exception of the dilution (1/1000).
6. Tyrosine hydroxylase (TH). Immunohistochemistry proceeds identically to parvalbumin with the exception of the primary antibody (Chemicon, 1/500).
Vasoactive Intestinal Peptide (VIP). Free‑floating sections are rinsed twice in phosphate‑buffered saline (PBS) at pH 7.4 for 5 min each and transferred to a buffered 0.6% hydrogen peroxide solution in order to block staining of endogenous peroxidases. Prior to the first antibody incubation, sections are placed in vehicle only for 20 min at room temperature. The vehicle (diluent) for all antibody incubations is 5% goat serum in PBS. Sections are then placed into a 1/2000 solution of primary antibody (Incstar) overnight at 4C. The next day, sections are transferred into a biotinylated goat anti‑rabbit immunoglobulin solution (Vector Laboratories) diluted 1/60 for 2 h at room temperature. After two washes in PBS, the sections are placed into ABC complex (Vector Laboratories) for 2 h at room temperature. The tissue is rinsed twice in PBS and twice in 50 mM Tris buffer (pH 7.6) and developed, dehydrated, counterstained, and coverslipped as with NF (see above).
GABA. The procedure for GABA staining is identical to that for VIP with the exception of the dilution of the primary antibody (Incstar), which is 1/333.
Somatostatin. The procedure for somatostatin (Chemicon) is identical to that for VIP. The dilution is 1/100.
Glial Fibrillary Acidic Protein. The procedure for GFAP staining is identical to that for VIP with the exception of the dilution of the primary antibody (Incstar), which is 1/25.
Potential Pitfalls and
Problems with Immunohistochemistry
Although antibody penetration is a potentially serious issue with immunohistochemistry, we have extensive experience with immunohistochemical procedures, and the antibodies we use have proved remarkably robust and reliable. We will pay careful attention to the quality of the tissue, and if necessary we will explore various methods to enhance antibody penetration, including the use of various detergents.
7. Financial Considerations
A. Additional Technical Support Needed
One full-time technician and
one half-time technician will carry out activities related to the
Neurohistology Core of the Neuroinformatics Program Project.
B. Additional Equipment Needed
An additional microtome is
needed for processing the large volume of tissue being generated by the
C. Additional Costs Relating to TravelTravel costs will be handled through the individual research projects.