Rb Anti-vGlut2 Preview

Rabbit anti-vGliuT2 (1:16k) in the LHA (approx. Level 28). Image was acquired with Volocity and assembled with Photomerge/PS (CS5). The original image was gamut and profile corrected, and cropped and rotated. This version was also, sharpened & brightened, converted to sRGB & reduced to 1024x1024 (From 3737 x 3737)

Immuno Test Results-11/16/12

The immuno came out great—basically all primaries tested worked best at the highest dilution tested. That is,

• 1:40k for Paul's MCH
• 1:16k for the rabbit anti-vGlutT2
• 1:5k for Zamir's MCH
• 1:10k worked well for the NEI 

To document the results we should image at least one section of each at as close to the same atlas level as possible.

Sections to image for Paul's MCH 
1:10k = S1 on the 10, 20 40k slide
1:20k = S4 on the 10, 20 40k slide
1:40k = S1 on the 40k slide (see below)

For Zamir's MCH, an image of S3 on the 1, 2.5, 5k slide will be enough

The NEI-labeled sections are generally too rostral or caudal for a direct comparison, but there is labeling in the LHA and ACB. Time permitting, it would be nice to image S1 (ACB) and S3 (LHA, Lvl 27).

The vGlutT2 needs to be imaged as a 1 panel z-stack at 20X to clearly distinguish labeled elements. The background is very high at 1:4k so image 8 and 16k only. 


I've captured the 1st image (Paul's MCH, 1:40k) using Volocity on the 428c Zeiss, leaving only 5 or so to go...

1024 X 1002 image of Paul's MCH (1:40k), acquired as a 10X,  5 X 5 composite with Volocity, and assembled using PS and Photomerge. 

LHA Imaging Protocol

After examining the test images and crunching the numbers, here's the image acquisition protocol for the LHA project:

Image Acquisition:

Acquire images at 10X, 2048x2048, 8 bit, spd 7, Avg 2, 5x4 tiles (15% overlap), with an optical slice thickness of 8.5 µm, which gives 4z in 25 µm sections or 6z in 40 µm sections.

z-Stack:
The slice thickness is based on a pinhole of 40.9 µm, which is approx. 1 A in the Far Red (FR), 1.08 A in the Red, 1.25 A in the Green and 1.66 A in the Blue channels.

Acquisition parameters are optimized across all 4 channels, even though we will rarely use the 4th channel. The alternative is optimizing each combination of tracers for the longest wavelength, which would mean that the x, y and z resolution (lateral resolution and z-step) of a channel would change depending on the combination of channels.

All of these parameters can be found in the 2nd and 3rd sheets of the Image Acquisition Google Document.

Imaging Strategy

The file size, acquisition time, and the number of immuno series to be imaged makes it impractical to image all available LHA sections (except perhaps in the 40 µm 1:6 series). Instead sections to be imaged will be based on Atlas levels prioritized by region of interest. For us (as opposed to UTEP), we will initially focus on the sections corresponding to Atlas level 26, then levels 24 and 25, followed by levels 29 and 30 and finishing with levels 23 and 32. These levels represent the iconic view of the LHA (26), the ACB-recipient LHA (24 and 25), levels central to Arshad's region of interest (29 and 30) and the rostral and caudal  LHA  ends (23 and 32; not quite the pole, but easier to find/identify). This strategy covers 8 of 11 levels of the LHA. Exactly how this translates into images acquired is difficult to estimate because it is likely that more than one image will to deal with plane-of-section issues.

Immuno Strategy

Assuming that the same antibody raised in a different species gives identical results, our goal is to prepare series for 11 different antibodies (plus AVP), replicating one or more labels in each brain to facilitate analysis of the patterns of expression across brain. Ensuring that each brain is in register with the others will require 5 brains of 4 series each, for a total of 20 immuno series, which works out to be 2 series each for 9 of the 11 series, and 1 series for the last 2 (which will likely be AChE and vGluT). Because we want a single-labeled series for each peptide, this also means that 11 of 20 will be single labeling (plus DAPI), and 9 of 20 can be double or triple labeled (plus DAPI).

Files Sizes & Acq Time

 Single Images:
2 Channels: 
10x, 2048, spd 7, Avg 2, 5x4x4x2 (25 µm) = 800 MB and 52 min/image
10x, 2048, spd 7, Avg 2, 5x4x6x2 (40 µm) = 1120 MB and 73 min/image

3 Channels:
10x, 2048, spd 7, Avg 2, 5x4x4x3 (25 µm) = 960 MB and 62 min/image
10x, 2048, spd 7, Avg 2, 5x4x6x3 (40 µm) = 1440 MB and 93 min/image

Level 26:
Let's assume we'll acquire 2 images (not counting the Nissl) to capture level 26 X 20 series to cover all the immuno = a library of 40 images.

If each image is 800 to 1400 MB, and takes somewhere between 52 to 93 minutes to acquire, imaging level 26 take anywhere from 35 to 62 hours and generate between 31 to 56 GB of images. 

All Images/Levels:
Let's assume it will take 12 images to cover the 8 levels of interest. That's 240 images to to cover all series. Using the same numbers for size and duration, it will take roughly 200 to 372 hours (8.5 to 15.5 days) and generate between 187 and 338 GB of images—not including the Nissls. Overall, these numbers aren't as bad as I expected, but clearly the sooner we start, the better off we'll be

 Next Steps

Three things are needed before we start generating data: 

• Work out the DAPI (or NeuroTrace) protocol and get accustomed to using it
• Generate the Nissls and figure out the correspondence between section number and atlas level
• Finalize our list of brains to use, 1º combinations (both which peptides go with each brain and which will be singles & which labels will be combined)

And only two things are needed until acquiring the data is all that remains:

• Optimize the remaining antibodies (5-HT, vGluT2, DYN, and any species variants such as sh ∝ GAD)
• Run the immuno...

ms ∝ GAD67 & ms ∝ VGlut2

In brief, all GAD67's worked great, and the 1:5k grouped worked the best (and should be used for all future GAD67 series). It could probably be diluted further, but isn't necessary (& therefore not worth the time to try it).

The VGlut2 on the other hand wasn't so clear cut. There was clear and obvious labeling in the MH at all dilutions, but it was difficult to say that differences in intensity in other brain regions were specific vGlut2 labeling. This may be due to the nature of vGlut2 labeling, and for this reason, tests should be run with the rb ∝ and/or gp ∝ VGlut2 before writing off the ms ∝ VGlut2.

Imaging Nomenclature

When working with 4-dimensional images, terms like 'image', 'slice' and 'section' become ambiguous, so here's a tentative nomenclature scheme for referring to an image and its parts. 

• Aperture: The pinhole opening that allows light to pass. Aperture can be measured in µm, which is constant, or Airy (A or AU), which varies with wavelength. Aperture also determines slice thickness. Note that the resolving power of a given aperture varies by wavelength, but slice thickness does not.
• Brain: All series volumes
• Channel: The part of the image containing the signal, or labeling. Each image can have multiple channels.
• File: The saved/stored/named components of an image
• Frame: One field-of-view (f-o-v). The most basic component image.
• Image: The assembled frames (Tiles + Stacks)
• Library: A collection of images from multiple brains
• Mosaic: An image composed of stitched tiles.
• Series: Set of brain sections collected at the same interval or frequency. Can also 
• Slice: One of two or more frames in register in z
• Stack: All slices in register in z
• Stitched Stack: A mosaic with x, y, and z components
• Section: A brain section.
• Slab: The z distance of a section that is imaged. Ideally, slab thickness = section thickness, but is usually less
• Tile: A component of a stitched image. Can be a frame or a stack.
• Volume: All sections of a series
• z-Step, or slice interval: The distance moved, in z before imaging the next slice. Note that Nyquist oversamping is 150%, meaning the actual number of slices is 1.5 x (slab thickness/slice thickness.

1024 vs 2048: x,y,z preview

512 x 512 image of NT labeled w/Alexa 488. Source is 10X, 2048x2048, single channel, and 2x2x5 tiles (20 component images, 4 MB/tile and 80MB for the image). Acquisition time was 15.49 s/ tile and 5m 15s for the image

512 x 512 image of NT labeled w/Alexa 488. Source is 20X, 1024x1024, single channel, and 4x4x16 tiles (256 component images x 1 MB/Tile = 256 MB for the image). Acquisition time was 7.75 s/ tile and 35+ m the image.

Larger verision of 10X, 2048: 900 x 897 (but compression = 7)

Larger verision of 20X, 1024: 900 x 897 (but compression = 7)

Clearly more detail available in the 20X image, but differences diminish at the perspective needed to view the whole image (original images are 3634 x 3594, which is approx. 12 x 12" at print resolution, and 38 x 38" at screen resolution!)

1024 vs 2048, Round 2

AVP 10X, 1024x1024: File = 512x512, 96dpi, sRGB, saved as JPEG (Q10)

AVP 10X, 2048x2048: File = 512x512, 96dpi, sRGB, saved as JPEG (Q10)

A single field of view of AVP labeling in the LHA was acquired at 10X for both, but a scan resolution of either 1024 x 1024 (top, left) or 2048 x 2048 (middle, right). Side-by-side comparison of the original, full-size images shows much greater (approx. 2X) effective resolution and detail in the 2048 x 2048 image. However, as the close ups show in the final two images, the benefit of the additional detail depends upon the output resolution and the size of the final image.

AVP 10X, 1024, cropped to 128 x 128 (= 2x original)

AVP 10X, 2048, cropped to 256 x 256 (= native resolution)

Methods: After acquisition, images were exported as TIFF, opened in PS and color converted (-->WG_RGB 1.8-->Adobe'98). A version of the 2048 image  reduced to 1024x1024 was created for comparison. Higher power comparison produced by cropping the originals to fixed size over a similar area. Posted images were additionally processed with Auto Contrast, reduced to 512x512 at 96 dpi, converted to sRGB and saved as hight quality (10) JPEG images.

Single Panel Comparison of 1A vs. 1.5A, 20X in Z

121027_005
20X,  1024x1024
1.01A, 7z at 1.8µm per slice, Gain 397

121027_004
20X, 1024x1024
1.50A, 6z at 2.6µm per slice, Gain 385

These two images are of the same field of view in of the rb anti VAS 1:10k series stained on 9/28/12. The purpose of the two images is a comparison of Airy units. Changing the aperture altered the light settings, so the gain and slice thickness was altered between images. All other variable remained the same.

Comparison Image Taken at 1.5A with 5 Z-slices

121027_002
10X 2048x2048, 1.47A
Saved as 936x936 jpeg, sRGB

This image was taken at 10X, with 5 z slices that were merged in a Maximum Intensity Projection. The image was acquired with 1.47A, sharpened and color enhanced in photoshop.

Single Panel Comparison of 1024 vs 2048 Acquisition

121023_001
10X, 1024x1024 resolution

121023_002
10X, 2048x2048 resolution

Both images are of the same field of view of tissue stained with rb∝AVP 1:10k (VAS, run on 9/21/12). All acquisition settings were kept constant (1.5 Airy), only the resolution was changed from 1024 to 2048. Images were edited in Photoshop using unsharpen mask and converted to sRGB jpeg. Image colors were highlighted using curves.

Elvanol Protocol

Elvanol (Polyvinyl Alcohol; PVA) is a polymer that penetrates the tissue and is used as a mounting media to firmly affix sections to the slide. To be used almost exclusively for mounting Nissls.

To prepare the PVA stock solution:

• Heat 1 L DI to near boiling (± 90º C) in the fume hood. Cover to prevent excessive evaporation. Use a thermometer to monitor temperature. If the DI boils, remove from heat and allow to cool.
• Remove from heat and gradually add 3.0 g PVA (Type II, Sigma #P1763) while stirring. Note: PVA dissolves slowly, taking as long as 30 min to go in solution.
• Add 1 g Na Azide* as a preservative. Note: This step is optional but especially useful if recycling/reusing the PVA.
• Let solution cool to RT, filter if needed and store at 4ºC

Mounting:

To mount, transfer sections to 6-well trays containing PVA, then mount as normal using PVA in place of buffer in the petri dish. No post-processing or rinsing needed. As for all other mounting, place mounted slides in racks and store racks in the drying oven until ready to stain.

Note: After mounting, PVA can be filtered and reused. Store filtered/recycled PVA in a separate stock bottle.


*Na Azide is toxic (works like CO, irreversibly binding to haemoglobin). Always were mask and gloves. It does not need to be weighed in the hood, but care should be taken to clean the balance and counter whenever and wherever Na Azide is used. Once in solution, gloves nor mask are needed while mounting.

Vasopressin 1024

121017_004 Cy2⦁rb∝VAS1:10k
1024x1024 per panel, 10x

AVP section taken from immuno series on 9/21/12 (slide 1, section 5). Image assembled with photomerge (CS5), converted to sRGB at 96 DPI. Saved as JPEG, quality set to 10 (of 12). Image size 768x623.
This image was taken primarily to compare with that taken at 2048x2048 per panel at 10X. This is the same tissue, all acquisition settings were kept the same with the exception of panel resolution.

Vasopressin Preview

Georgina's AVP (002) 10X Composite photo, assembled w/Photomerge (CS5). Image is 2048 x 1614 or 21.3 x 16.8" at 96 dpi, converted to sRGB and saved as JPEG. Quality set to 10 (of 12). Size as uploaded = 1.2 MB.

The cropped version of the full resolution is 96.4 X 75.9" at 72 dpi and 114 MB when flattened...

NT & NOS Double

Labeling with antibodies raised against FITC allows amplification of both signals when double labeling. Cy 2⦁ms∝FITC (Jackson) and Dylight 488⦁gt∝FITC (Rockland) have also been tested. All work well (A488 = Cy2 > D488) providing approximately the same degree of signal amplification as Streptavidin.

Double labeling for mouse∝NOS1 (red) and rabbit∝Neurotensin (NT; green), labeled with Cy3⦁Streptavidin and Alexa 488⦁rb∝FITC (Invitrogen), respectively. Component channel images below.

Note that image was acquired as a 4 X 5 2-channel mosaic. Component tiles were exported for each channel. Images of each channel (below) were assembled separately w/ Photomerge--which put the images together slightly differently. Mis-registration can be clearly seen along ventral-lateral surface.

Individual red (NOS) and green (NT) channels. Both received minimum crop and autotone adjustment only. Merged image above cropped and rotated, curve adjustments and unsharp mask applied.

Slide Subbing

Note: This protocol is from Jackson Bittencourt/Paul Sawchenko (1989), and is an abbreviated version of the lab protocol (from Simmons et al., 1989). My version was originally posted Oct. 3, 2012 and revised Feb. 12, 2013

1. Dump new slides into Acid Alcohol (70% EtOH + 1% HCl)
2. Hand wash* and transfer to distilled (DI)
3. Rack slides leaving one blank space at either end.
4. Soak the racked slides in DI + Alconox* for 30 min or more
5. Rinse out Alconox in DI for 1 - 2h, and leave soaking in DI overnight if possible
6. To preparing the subbing solution, use:

• 500 ml* DI in a 1000 ml beaker
• a funnel (preferably glass, but must be clean)
• ring stand
• filter paper
• 5.0 g gelatin (Knox Gelatin*, available at most grocery stores)
• 0.5 g Chromium Potassium Sulfate (On the shelf with the gelatin)

      5.  Heat water to 70 - 80º C (i.e., do not boil)

      6. Add gelatin and, after gelatin dissolves, add Chromium Potassium Sulfate

      7. Let cool to 40º - 60º C* and filter into glass Wheaton staining dish 

      8. Soak slides/racks for about 2 min each

      9. Put slide racks on tray, tipped to drain with frosting pointing down. Cover with

          lab diaper to protect from dust

    10. Move tray(s) to oven (37 - 40º C) overnight.

    11. Remove from oven & let cool to room temperature before putting away.

    12. Store slides in clean, dust free boxes. Wrap in plastic wrap for long term storage

*Notes:

1. New slides are often obviously dirty so the acid alcohol wash is an annoying but necessary step
2. Hand washing is a quick swipe front and back with a 4 x 4 gauze sponge, or similar cheap, sturdy, disposable and lint-free cloth.
3. Alconox. Use 7.5 g/L and make about 6 L (45 g) for 10 large racks. More soluble and effective in hot water. Recommend heating 3 L DI to 40 - 60º C, adding 45 g Alconox while stirring, then at to 3 L DI (RT), rather than using hot tap water
4. Subbing: 500 ml is barely enough to cover a rack in a large dish and gelatin is cheap so make plenty. Time can be save by subbing 2 racks in parallel. For 2 dishes make 1.5 L subbing sol'n.
5. Gelatin: The  Knox packs contain approx. 7.2 g gelatin,which means 3 packs are needed to make 1.5 L
6. Subbing works better with hot gelatin but is simply too hot to handle above 60º
7. Stored slides should be labelled as "Sub'd" and dated but last indefinitely if wrapped (i.e., protected from dust and moisture)

The secrets to subbing are keeping the slides and dishes clean, avoiding bubbles, and don't over do it. To keep the slides clean, 1) wear gloves (hands are oily, if not dirty),  2) avoid lint. Don't use paper towels for anything. Do use Kimwipes and other papers stated to be lint free. Set slides on the absorbent side of the lab diaper, but cover with the plastic side facing the slide, and 3) keep the slides either submerged or covered (even in the oven)  To keep the dishes clean, wash them immediately (because obviously dried gelatin doesn't come off glass easily). Because dust is the biggest enemy, the Wheaton dishes should be dried also and wrapped in plastic wrap for long-term storage. Use lint-free Kimwipes, not paper towels. Bubbles are a problem because they dry as raised rings of gelatin, potentially causing slides to stick together and always rendering the slides unusable. You always get bubbles when filling the Wheaton dish. Use a Kimwipe to remove bubbles by blotting them away or at least moving them to the sides of the dish and out of the way. When removing racks from the subbing solution, drain well and inspect each rack of slides making sure that there is no liquid remaining in the gaps between slides. 

Finally, figure out how many slides can be subbed and do that many or less. The best policy is to use entire boxes. Open, partial boxes should be discarded to avoid confusion and costly mistakes. Note that slides come 72/box and 20 boxes/case which means 1440 slides in a case. Currently we have enough racks to do about 15 boxes (> 1000 slides) in a run, but only one tray that fits the oven. The tray holds 8 large racks, which combined could take 384 slides but only 5 boxes (360 slides). Buy one more tray and the max run increases to 10 boxes, which is probably the practical capacity of the oven. The actual run should be limited to the number of slides that can be subbed without compromising on any of the steps in the protocol.

      

Large, 50-slide metal slide rack. Similar to 25-slide rack

Wheaton Staining dish to fit large slide rack

TRIS buffers & Stocks

To make a 10X Stock for 50 mM Tris Buffered Saline (TBS), pH 7.2,

Add 1 L blue cap DI to:

• 70.2 g Tris HCl
•   6.7 g Tris Base
•    90 g NaCl

To make 1 L TBS working solution, add 100 ml TBS 7.2 Stock to 900 ml blue cap DI

Alkaline phosphate substrates require high concentraion, high pH buffers. Primarily for this reason we also occasionally make 10X 50 mM TBS, pH 8.0 and 5X 200 mM TBS, pH 8.5.

To make the 50 mM TBS, pH 8.0 10X Stock,

Add 1 L blue cap DI to:

•  44.4 g Tris HCl
•  26.5 g Tris Base
•     90 g NaCl

To make the 200 mM TBS, pH 8.5 5X Stock,

Add 1 L blue cap DI to:

•  44.4 g Tris HCl
•  87.2 g Tris Base
•     45 g NaCl

To make 1 L working solution from a 5X Stock, add 200 ml of the TBS 8.5 Stock to 800 ml bcDI

To make Tris buffers at other pH's, Molarities or for use at temperatures significantly colder ((4ºC) or hotter ( (37º) than RT, refer to  Sigma's Tris Mixing Table (pdf)

Imaging Records

When imaging, we need to record enough information to find the images we need, know what we're looking at when viewing an image, be able to reproduce the image or acquire other images in the same way, and of course to describe all relevant details in a manuscript's  methods section. Below is a detailed list of information to be recorded with explanation of the entry, which we may want to use to create a printable template or a Google document form.

Layout: Organizing the record into sections (Header, Source, Content, Acquisition, and File Info) will help ensure all info is recorded and make it easier to find things and compare settings across images and session.

Session Header 
This section contains information that applies to the entire session. Part is entered at the start of the session, and the part at the end of the session.

• Date
• Microscope
• What is being imaged (immuno and brain region)
• Total # of images acquired
• Range of file #'s of related images (e.g., same brain region, but different objective or acquisition parameters)

Image Source 
Imaging source information is recorded after the header and for each image as needed. That is, if the imaging session is organized the Image Source info can organize the record into sections, making it easy to add the summary info at the end of the session.

• Experiment (the immuno run and date; should be the 1st two lines on the slide)
• Brain and series (e.g., G1110c)
• Section # in either/both relative o/andr absolute notation. Relative notation refers to slide and section (e.g., 2-3 for first slide, third section). Absolute notation refers to the nth section of the entire series (e.g., S23, which is probably the 5th section on the 4th slide (4-5)). Relative numbering makes it easier to find the slide. Absolute numbering helps keep slides in register across series. For the LHA immuno series, the absolute section numbers will be noted on the slide

Image Content

• Image number (Starting with Image 01. Number consecutively for all images acquired in the session)
• Immuno and labeling (What's labeled and what it's labeled with; e.g., ms∝NOS (Cy2), rb∝NT (Cy3))
• Brain region imaged
• Objective (10X, 20X, etc.)

Note that this information can be combined into a sentence, e.g., 

Image 01 Tiled, 10X image of NOS/NT (Cy2/Cy3) in the LHA

Image Acquisition

• Channels. For each channel note

• Laser line(s) or Fluorescence Filter(s). 

• Laser Power for each line

• Scan Speed (6 - 8, usually 7)
• Averaging (1, 2, or 4)
• Aperture in Aiery (e.g., 1 or 1.51, etc.), in µm and slice thickness (in µm).
• Gain
• Offset
• Tiles (e.g., 3 X 4)
• Slices (number and slice thickness in µm)
• Acquisition Time (per tile and for entire image)

File Info

• Dimensions (generally 1024X1024 per channel)
• Bit depth (8 bit for now)
• File name. File naming should be date_image #_image content e.g.,

121001_01_LHA_Cy2NOS Cy3NT.cvi

or

121001_01_LHA_Cy2NOS Cy3NT_10X_9 tile_3 z.cvi

• File name of export. All files should also be exported as an OME.tif. The file name will generally be the same as above, but appended with the new file type
• Storage Location. Initially files will be stored on the same computer where acquired but at the end of the session a copy of all files should be made and transferred to a yet to be determined storage location.

Cy 3 GAD Mosaic

Immuno: rb∝GAD (1:40), labeled w/ Cy3⦁SAvidin (1:760, 90 min)
Tissue: 9/21/12, G1110C
Image: 10X, 9 panels, 1024x1024, 8 bit, 1 channel = 2747 X 2745 (cropped)=38x38" @ 72 dpi
Acquistion: 555 line, 2.0%, 1.58 Airy, Spd 7, Avg 2, Gain 559, Offset 0; 7.75 sec/panel
Tile: 15% overlap; Rotation not corrected (Zen could not stitch image)
Stitching: PS Photomerge, Auto mode
Processing: In PS, Assign & Convert to profile, Flatten and Crop. Colorized with Curves; Unsharp Mask: (Split: 64% Lighten, 31% Darken); Reduce size (from 9x9 to 6x6), convert to sRGB, save as JPEG
File Name: GAD 40k 10x 9 tile sm sRGB.jpg
Original File: 120921_Cy3GADA_40k_10X_9 tile_m0(...m8).tif

Random image of Cy 3 GAD in CP & overlying CTX. Captured for simple (fast imaging, limited tiles) source images to stitch. Conclusion: Zeiss Zen is capable of stitching images with 1 or fewer tiles...

Alexa 488⦁rb∝FITC

Immuno: rb∝Neurotensin (1:8k) labeled with Alexa 488⦁rb∝FITC (1:2K, Invitrogen)—to be used in double labeled series, in combination with a streptavidin conjugate. We also have Cy 2⦁ms∝FITC (1:2k, Jackson), which works just as good as the Alexa, and Dylight 488⦁gt∝FITC (Rockland), that's being tested now.
Image: 20 panels at 10x, 1024x1024 (x20), 8-bit
Acquisition: 488 line at 1.8%, 1.5 Airy, Gain 569, Offset 0, Speed 6, Avg 4 (31 sec/panel; 10m 30 sec total)
Tiling: 20% overlap; Rotation Correction 0.8087º (calibrated from Zeiss-provided macro from within Zen)--& the Zeiss software still wouldn't stitch
Stitching: PS 5 photomerge, auto mode.
Image Processing: In PS Assigned profile Prophoto and converted to Adobe '98. Flattened, Cropped and resized. Colorized with Curves. Sharpened with Unsharp Mask. Reduced size, converted to sRGB and saved a copy as a JPEG

Conclusion: Great staining, amplifies as much or more than streptavidin. However, 10x 1.5 Airy clearly doesn't capture all the data (when viewed at actual size of the original which is 14 x 11" at 300 dpi) and, unlike the NOS, would probably benefit greatly from acquiring in Z...

Neurotensin (1:8k) labeled with Alexa 488⦁rb∝FITC (1:2K)—to be used in double labeled series, in combination with a streptavidin conjugate.

DAPI Protocol

1. Order DAPI (4',6-Diamidino-2-Phenylindole, Dihydrochloride). Available from Invitrogen as either D1306 (10 mg for $85.68) or D21490 (10 mg, ultrapure for $126.63)
2. Make stock solution of 5 mg/ml by adding 2 ml DI to one 10 mg vial. Stock solution = 14.3 mM. Note that sonication may be required to get it into solution
3. Stain. Incubate tissue in 300 nM DAPI for 1 - 5 min.
4. Rinse, coverslip, mount.

Note:

• 300 nM is a 1:47667 dilution of 14. 3 mM

which means

• 1 µl DAPI stock makes 47.7 ml DAPI working

which means

•  2 ml of stock will make 95.33 liters of DAPI stain

Dilution of Streptavidin Conjugates

Working Dilution of Streptavidin conjugates from Jackson Immuno is now standardized to a working concentration of 2 µg/ml, which is the upper end of Jackson's recommended 1 - 2 µg/ml.

Calculations are based on:

1. Each vial of streptavidin contains 1.0 mg, regardless of conjugate
2. Every conjugate is a different concentration (HRP at 1.0, Cy 2 at 1.2 and Cy 3/Cy 5 at 1.8 mg/ml)
3. Each concentration is reconstituted with a different volume of DI (1 ml, 950 µl and 650 µl, for HRP, Cy2 and Cy 3/Cy 5, respectively)

Calculations for 2 µg/ml:

 HRP⦁Streptavidin (1:500)

HRP⦁Streptavidin comes as 1.0 mg, rehydrated with 1.0 ml DI = 1 µg/µl. Therefore 2 µl's/ml are needed for a working concentration of 2 µg/ml. 2 µl's/ml = 1:500, or 20 µl per aliquot makes 10 ml at 1:500. 1 ml theoretically makes 50 aliquots at 20 µl per aliquot

 Cy 2⦁Streptavidin (1:526)

 Cy 2⦁Streptavidin comes as 1.0 mg, at a concentration of 1.2 mg/ml, and is rehydrated with 950 µl DI. 1 mg in 950 µl = 1.05 µg/µl and 2 µg = 1.9 µl. Therefore 1.9 µl's/ml are needed for a working concentration of 2 µg/ml, which would be 19 µl in 10 ml, which is a dilution of 1:526. Note that again 950 µl theoretically makes 50 aliquots at 19 µl per aliquot.

 Cy 3⦁Streptavidin (1:769)

 Cy 3⦁Streptavidin comes as 1.0 mg, at a concentration of 1.8 mg/ml, and is rehydrated with 650 µl DI. 1 mg in 650 µl = 1.54 µg/µl and 2 µg = 1.3 µl. Therefore 1.3 µl's/ml are needed for a working concentration of 2 µg/ml or 13 µl per aliquot for 10 ml, which is a final dilution of 1:769. Here, too, 650 µl theoretically makes 50 aliquots at 13 µl per aliquot

 Cy 5⦁Streptavidin and Alexa 647⦁Streptavidin (1:769)

 Cy 5⦁Streptavidin also comes as 1.0 mg, at a concentration of 1.8 mg/ml, and is rehydrated with 650 µl DI. 1 mg in 650 µl = 1.54 µg/µl and 2 µg = 1.3 µl. Therefore 1.3 µl's/ml are needed for a working concentration of 2 µg/ml or 13 µl per aliquot for 10 ml, which is a final dilution of 1:769. Here, too, 650 µl theoretically makes 50 aliquots at 13 µl per aliquot

IHC Test Results: 9/21/12

9/21/2012 Immuno Results (Preliminary)

General: Tissue in tatters, few sections per group, & many sections missing, but everything appeared to work and the sections were mounted in great anatomical register

Short vs Long: Seems clear that the short runs (2 hr in 2º, 90 min in SAvidin) worked as well as the long run (3+ hr in 2º, 2 hr in SAvidin). For the vGAT "blind control", where run conditions were not noted on either slide, I couldn't really tell the difference.

rb ∝ GAD (1: 20, 30 & 40k): All dilutions worked, but (I think) 1:20k was best--more because labeling was good and background was low, so there is little cost to going with 1:20k, rather than some particular gain

rb ∝ DYN (1: 5, 10 & 15k): Could see well-labeled fibers at all dilutions, but no clear concentration of fibers in a recognizable terminal field. Could see lots of labeled cells, somewhat concentration dependent so am not absolutely sure that this is specific labeling. To accurately assess, comparison with DAB series needed to identify the most prominent terminal fields, and the best sections/levels. If not present in these test sections, then the test series will need to be run again

rb ∝ vGAT (1: 2.5, 5 & 10k): Labeling saturated making it difficult to distinguish individual components at 1:2.5k, and it looked as though labeling may have fallen off a bit at 1:10k. Labeling at 1:5k looked great (although I'd bet that 1:7.5k would be optimal). As noted above, labeling group was not recorded on either slide.

Alexa 488●rb ∝ FITC (1: 750, 500 & 2k): Note that the original plan called for 1: 750, 1500 (= 1.5k) & 2k, so only 750 and 2k dilutions mounted. Both worked/looked great. However, at 1:2k there appeared to be some fibers better labeled than others. Did not make comparisons needed to tell whether this was due to concentration, or fiber size (or similar). To make best determination, further comparison of 1:750 & 2k groups needed, as well as comparison of ∝ FITC with SAvidin-labeled NT. However, the 1º has been in solution at 4ºC since July 2010, so I took my best guess and aliquoted the 1º in order to store it at -70ºC.  Aliquots contain 15 µl, which makes 15 ml at 1:1k, or 30 ml at 1:2k 

TRIS-Buffered Glycerol, pH 7.2

Recipe: Tris-Buffered Glycerol (TBG), 5% n-Propyl Gallate (nPG), pH 7.2

To make 10 ml TBG 7.2:
● Add 0.5 g nPG to 1 ml 2M Tris, pH 7.2*
● Add nPG/Tris to 9 ml glycerol.
● Mix well, aliquot and store at -20º C

For 50 ml TBG 7.2:

● Add 2.5 g nPG to 5 ml 2M Tris, pH 7.2*

● Add nPG/Tris to 45 ml glycerol.

● Mix well, aliquot and store at -20º C

Note:

1. 2M Tris = 1.6 g Tris HCl + 0.134 Tris Base in DI (blue-cap or better). Note that a calomel electrode is needed to accurately measure Tris pH
2. nPG goes readily into solution at about 60ºC. Use a water bath heated to 65 - 70ºC to allow time to vortex nPG solution and add to glycerol without precipitating out of solution (and have glycerol measured and ready...)
3. 500 µl aliquots are enough to coverslip 6 to 8 slides depending on coverslip size, and yield at best 20 aliquots from 10 ml. Scintillation vials and 50 ml centrifuge tubes work well for making larger volumes
4. nPG is an antifade agent that slows bleaching of cyanine and alexa dyes, but can be omitted if desired.
5. Recipe was adapted from Confocal Microscopy for Biologists (Hibbs, A.R., 2004; Springer).

Results & Rationale
Originally tested 8/16/2012 as a means of limiting the excitability of Fluorogold (FG) by wavelengths longer than UV, or at least < 488 nm to keep it out of the green (and red channels). 

Two TBG mounting media at pH 7.2 and 7.6 were compared to the normal bicarbonate (KHCO3)-buffered glycerol (BBG, pH 8.6), and tested on FG-injected as well as green- or red-labeled peptide IHC.

Both TBG media both reduce the number of FG-labeled neurons visible in the green and red channels compared to BBG without attenuating the FG signal or interfering with normal peptide IHC. However, TBG, 7.2 was clearly more effective, eliminating all neuronal labeling in both channels. At the injection site, FG labeling was clearly visible in the green and red channels when mounted w/either BBG or TBG 7.6, but faintly visible in the green channel only w/TBG 7.2. 

New PBS-PFA Recipe

The PBS-PFA tested on 9/4/2012 seemed to work fine, but for the future we are changing to a more common variant of Sorensen's phosphate buffer, based on recipes from  UC Berkley Biological Imaging Resources and Wikipedia—Phosphate Buffered Saline, and adapted for use with anhydrous Na2HPO4 (Berkley used 7●H2O, and Wikipedia 2●H2O)

PBS, pH 7.4: 

For 1 liter of 20X Stock use:

KCl                                         4 g

NaCl                                   160 g

Na2HPO4 (anhydrous)          22.9  g

(Sodium Phosphate Dibasic)

KH2PO4                                  4 g

(Potassium Phosphate Monobasic)

For 1 liter of PBS-Working, add 50 ml 20x stock to 950 ml DI

4% PFA in PBS, pH 7.4:

• For 1 liter of 4% Paraformaldehyde (PFA) in PBS, add 40 g PFA per liter of working PBS 
• For 4 liters of 4% PFA in PBS, add 200 ml 20x PBS Stock to 3800 ml DI.
• Heat to 60 - 70º C (Do Not Exceed 70ºC) and add 160 g PFA. Add NaOH (0.5 g/l) if needed to get PFA in solution. 
• Filter the PFA, with coffee filter, when cool enough to touch
• Move filtered PFA to 4º C overnight, or use salted-ice bath if perfusing the same day 
• Measure pH of cold PFA (4 - 8º C) and adjust pH to 7.4 with a few drops of HCl (12 N) or NaOH (1 to 5 N) if needed. 
• Prepare Sucrose Postfix (10 to 20% Sucrose in cold, pH'd 4% PFA). 
• Postfix for one animal = 11.5 g sucrose in 75 ml PFA (= 15%). For 3 animals = 35 g sucrose in 225 ml PFA; and postfix for 4 animals = 46 g sucrose in 300 ml PFA.

Perfusion Protocol:

Perfuse animals with normal saline (9g NaCl/liter) for about 5 min at a pump setting of 1.1 (= 40 ml/min) and then switch to PFA. When rigor sets in (or about a minute, whichever comes first), reduce pump speed to 0.8 (= 25 ml/min) for 20 min. Total volume of PFA perfused should be just over 500 ml. Remove the brain and place in cold sucrose postfix for 18 to 24 hr.

Protocol- Immuno Log Entry

Date                Antibody Used (dilution; tissue)

• Transfer tissue to 24 well nets
• Rinse out cryo w/ 50mMTBS7.2 X # rinses, time of each rinse
• LTBS= ?mL 50mMTBS7.2 + ?µl 25%T + ?mL NDS
• 1˚-
• 1 aliquot= ?µl 
• 1:?k= take ?µl 
• add ?ml LTBS= ?ml 1:?k
• use ?mL for 1:?k
• 1:?k= take ?mL 1:?k
• add ?mL LTBS= ?ml 1:?k
• use ?mL for 1:?k
• 1:?k= take ?mL 1:?k
• add ?mL LTBS= ?mL 1:?k
• Pre-incubate/Preabsorb for ? min
• Transfer to 1˚ -> 4˚ @ time

Date

• Rinse out 1˚ @ time
• total time in 1˚
• 50mMTBS7.2 X # rinses, time of each rinse
• Sort sections for secondary reaction
• LTBS= ?mL 50mMTBS7.2 + ?µl 25%T + ?mL NDS
• 2˚-
• name the secondary being used at what concentration
• calculate the dilution of the secondary
• Pre-incubate/preabsorb for ? min
• Transfer to 2˚ for ? time
• Rinse out 2˚ at @ time
• total time in 2˚
• 50mMTBS X # rinses, time of each rinse
• 3˚-
• name the fluor label being used at what concentration
• calculate the dilution of the label in 50mMTBS7.2
• Transfer to 3˚ at ? time
• Rinse out 3˚ at ? time
• total time in 3˚
• 50mMTBS X # rinses, time of each rinse
• Transfer sections to 6-well trays @ time to await mounting

Protocol- Anti-FITC Immuno Template

Schedule for FITC Immuno Round Completion:

Single labeling:
Friday- Sections into Primary (5x5min rinses, 30min pre-incubation/pre-absorption)

Monday- FITC(DK)anti(RB/MS) 1:500 in LTBS (5x5min rinses, 30min pre-incubation/pre-absorbtion,  
             2-3hrs in 2˚
             Alexa488(RB)anti(FITC) 1:500-2k? in TBS ~36hrs

Wednesday- Cy2(MS)anti(FITC) (1:1-2k) in TBS ~2-3hrs

Double labeling: 
Friday- Sections into Primary (5x5min rinses, 30min pre-incubation/pre-absorption)

Monday- Biot(DK)anti(RB) 1:500 in LTBS (5x5min rinses, 30min pre-incubation/pre-absorbtion, 2hr)
            FITC(DK)anti(MS) 1:500 in LTBS (5x5min rinses, 30min pre-incubation/pre-absorbtion, 2hr)
            Cy2(MS)anti(FITC) 1:500-2k in TBS ~36hrs

Wednesday- Cy3SAv 1:500 in TBS ~2hrs
                 

OR

Friday- Sections into Primary (5x5min rinses, 30min pre-incubation/pre-absorption)

Monday- Biot(DK)anti(MS) 1:500 in LTBS (5x5min rinses, 30min pre-incubation/pre-absorbtion, 2hr)                   
               FITC(DK)anti(RB) 1:500 in LTBS (5x5min rinses, 30min pre-incubation/pre-absorbtion, 2hr)
               Alexa 488(RB)anti(FITC) 1:500 in TBS ~36hr

Wednesday- Cy3SAv 1:500 ~2hrs
               (5x5min rinses)
                 

Recipe- Perfusate

Tested on 9/4/2012 (pH when 4˚ ~7.5)

To make 1 Liter PBS:

1Liter Blue Cap DI Water
NaCl 6.8g
Na2HPO4 1.5g
NaH2PO4 0.43g

add 4% Paraformaldehyde (40g/L)