Adding links to video tutorials from Leica.

data/videos.csv

@@ -6,4 +6,10 @@ XTRegisterSameChannel - SimpleITK Imaris Python Extension,https://www.youtube.co
 Coating slide with chrome alum-gelatin adhesive,https://www.youtube.com/watch?v=ksNR3gsl5rg,"Coat a glass slide by adding 15 &mu;L chrome alum-gelatin to one side. Spread evenly using the edge of a cover slip or a separate glass slide via the blood smear technique. Note, it may require multiple spreading passages with the slide to ensure even coating of chrome alum-gelatin without streaks. If chrome alum-gelatin pools on the slide, aspirate excess fluid and respread as these accumulations can result in autofluorescence artifacts during image acquisition. Be mindful of stated expiration date on chrome alum-gelatin.<br><br>For best results, prepare chrome alum-gelatin coated slides freshly (i.e., the day of tissue sectioning) or no more than 7 days prior to tissue sectioning. If chrome alum-gelatin coated slides are prepared ahead of sectioning, store at room temperature (do not freeze as this will compromise the adhesive properties).<br><br>Dry coated slide in oven at 60°C for 1 hour.<br><br>We purchase chrome alum-gelatin from Newcomer Supply, but recipes are available online if this item does not ship to your country (e.g. [this recipe](https://www.laboratorynotes.com/preparation-of-chrome-alum-containing-gelatin-solution-for-preparation-of-coated-slides-for-histological-tissue-sections/))",tutorial,2023,8,22
 Mounting tissue and applying coverslip,https://www.youtube.com/watch?v=tqHlrSmsG_8,Remove as much PBS as possible without drying out tissues. Quickly add the minimum amount of Fluoromount-G mounting medium necessary to completely cover each tissue section. Gently cover with a coverslip. We typically use 10 – 40 &mu;L per tissue section. Ensure there are no bubbles on or near tissue.,tutorial,2023,8,22
 Coverslip removal after image acquisition,https://www.youtube.com/watch?v=gXN19na8I6Y,"After the slide (containing coverslip) has been soaked in 1X PBS pH 7.4 to allow coverslip to loosen (typically 30-60 minutes), carefully remove slide from container. Ideally coverslip will have fallen off naturally. If coverslip is loose but still attached, gently lift a corner of the coverslip with a pair of fine forceps or razor blade.<br><br>We typically wait until coverslip detaches from slide naturally or, at minimum, is moving freely on the slide under the force of gravity. Attempting to remove coverslip before it has sufficiently loosened may result in tissue deformation and/or loss.",tutorial,2023,8,24
-Making 1 mg/mL of LiBH<sub>4</sub>,https://www.youtube.com/watch?v=-MH_aLEMyF8,"Dissolve LiBH<sub>4</sub> (STREM Chemicals, cat no. 93-0397; purchase in 1 g aliquots) into diH<sub>2</sub>O (Quality Biological, cat no. 351-029-101) and allow to sit at RT for 10 minutes until the formation of bubbles occurs (not shown in video). Pass the solution through a 0.22 &mu;m syringe filter (Millipore Sigma, cat no. SLGSM33SS) attached to a 20 mL syringe (EXEL Int., cat no. 26280) to remove any impurities prior to use (not shown).<br><br>Always prepare LiBH<sub>4</sub> solution in a chemical fume hood with appropriate personal protective equipment (PPE). Mixing LiBH<sub>4</sub> with diH<sub>2</sub>O will generate hydrogen gas which is highly flammable. For this reason, we always work with small amounts of LiBH<sub>4</sub> relative to the amount of water initially added. We do not mix >10 mg of LiBH<sub>4</sub> with diH<sub>2</sub>O. This amount can be quickly diluted to 1 mg/mL with volumes handled by standard serological pipets/pipet-aids. Always wrap LiBH<sub>4</sub> stock with parafilm (suggest Bemis Company Inc., cat no. S37440) and store in the presence of desiccant. We recommend replacing LiBH<sub>4</sub> every 4 weeks as repeated exposure to moisture/air has been observed to reduce fluorophore inactivation efficacy. Be sure to follow institutional guidelines for proper disposal of hazardous chemicals.<br><br>The formation of bubbles indicates the solution is ready for filtration and subsequent use. For best results, use solution immediately after filtration. We have noted effective fluorophore inactivation up to 4 hours after initial dissolution of LiBH<sub>4</sub>.",tutorial,2023,8,23
+Making 1 mg/mL of LiBH<sub>4</sub>,https://www.youtube.com/watch?v=-MH_aLEMyF8,"Dissolve LiBH<sub>4</sub> (STREM Chemicals, cat no. 93-0397; purchase in 1 g aliquots) into diH<sub>2</sub>O (Quality Biological, cat no. 351-029-101) and allow to sit at RT for 10 minutes until the formation of bubbles occurs (not shown in video). Pass the solution through a 0.22 &mu;m syringe filter (Millipore Sigma, cat no. SLGSM33SS) attached to a 20 mL syringe (EXEL Int., cat no. 26280) to remove any impurities prior to use (not shown).<br><br>Always prepare LiBH<sub>4</sub> solution in a chemical fume hood with appropriate personal protective equipment (PPE). Mixing LiBH<sub>4</sub> with diH<sub>2</sub>O will generate hydrogen gas which is highly flammable. For this reason, we always work with small amounts of LiBH<sub>4</sub> relative to the amount of water initially added. We do not mix >10 mg of LiBH<sub>4</sub> with diH<sub>2</sub>O. This amount can be quickly diluted to 1 mg/mL with volumes handled by standard serological pipets/pipet-aids. Always wrap LiBH<sub>4</sub> stock with parafilm (suggest Bemis Company Inc., cat no. S37440) and store in the presence of desiccant. We recommend replacing LiBH<sub>4</sub> every 4 weeks as repeated exposure to moisture/air has been observed to reduce fluorophore inactivation efficacy. Be sure to follow institutional guidelines for proper disposal of hazardous chemicals.<br><br>The formation of bubbles indicates the solution is ready for filtration and subsequent use. For best results, use solution immediately after filtration. We have noted effective fluorophore inactivation up to 4 hours after initial dissolution of LiBH<sub>4</sub>.",tutorial,2023,8,23
+Leica LAS X Navigator: Focus Map,https://www.youtube.com/watch?v=SbDDzo8dPZo,"This is a video tutorial on how to set up a focus map in LAS X Navigator (v. 3.6.0 Widefield), which is recommended for tissue sections or samples that are continuous.",tutorial,2020,6,9
+Leica LAS X: Hardware Autofocus (AFC),https://www.youtube.com/watch?v=3uq42XefHhY,"How to use Adaptive Focus Control (AFC) in your Navigator experiments, if your DMi8 microscope is equipped with AFC.",tutorial,2020,6,9
+Leica LAS X Linked Shading: Fluorescence,https://www.youtube.com/watch?v=9l_DmMmAbHk,A video tutorial on how to do Linked Shading (shading correction) in LAS X 3.6 (widefield) with fluorescent images on a monochrome camera.,tutorial,2020,6,9
+Leica LAS X THUNDER Tutorial,https://www.youtube.com/watch?v=36HqrFyy0fQ,"This video shows how to ""THUNDER"" images in LAS X on-the-fly or post-acquisition with the default settings. (v. 3.7.1 - Widefield). This is only available on THUNDER Imagers.",tutorial,2020,6,9
+Leica LAS X Software Experiment Setup: Z-stack,https://www.youtube.com/watch?v=dTMF01cO7lI,"How to set up a Z-stack in LAS X v 3.3 (widefield).
+Note: the example in this video was for a data set that was intended for deconvolution, so it goes slightly beyond the focus on either end. The usual recommendation is to stay mostly in focus for all of your Z planes, so that subsequent 2D projections (such as a maximum intensity projection or extended depth of focus projection) will be clearer.",tutorial,2020,6,9