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.