This work is a part of research program P3-0327 and research project J3-7177, founded by the Slovenian Research Foundation.

All methods described here have been approved by the National Medical Ethics Committee on 19 April 2016 (No. 0120&#8211;204/2016&#8211;2).
1. Set-up of the Microscope Recording System
<ol>
	<li>Turn off the heating plate of the microscope.</li>
	<li>Before any treatment, take a snapshot of a blastocyst under a camera-equipped inverted microscope. Remember to note the magnification at which the blastocyst was recorded.</li>
</ol>
2. Selection and Prepreparation of Blastocysts for Vitrification
<ol>
	<li>Select only fully expanded day-5 or day-6 blastocysts for cryopreservation with at least a few inner cell mass (ICM) cells and cohesive trophectoderm.</li>
	<li>For laser-treated blastocysts with a collapsed blastocoel, subject a blastocyst to a short laser pulse (0.4 - 0.7 ms) with a hole diameter ranging from 4.3 to 9.4 &#181;m, tangentially directed at the zona pellucida and at a junction of two adjacent trophectodermal cells. Allow the blastocyst to fully or partially collapse for 5 min.</li>
	<li>For untreated blastocysts with an intact blastocoel, perform no particular treatment before vitrification. 
	Note: These blastocysts are considered intact.</li>
</ol>
3. Preparation of Cryoprotectant and a Petri Dish for the Equilibration Phase
<ol>
	<li>Use pithe pipette for oocyte denudation (diameter 135 &#181;m) to aseptically fill equilibration media (M-199 HEPES-buffered medium with 7.5% DMSO, 7.5% ethylene glycol, 20% DSS) into the holes of the microdroplet area of a 9-well Petri dish specially designed for time-lapse microscopy and recording. 
	Note: Oocyte denudation is a process of removing cumulus cells surrounding the oocyte. The use of a pipette for oocyte denudation will help to avoid the creation of air bubbles.</li>
	<li>Overlay the microdroplet area with 30 &#181;L of equilibration media.</li>
</ol>
4. Transfer of the Blastocyst in Equilibration Solution
<ol>
	<li>Submerge a laser-treated or intact blastocyst into the equilibration medium to the bottom of the microdroplet area of the microdroplet dish for 10 min at room temperature (equilibration phase).</li>
	<li>Start a 10-min countdown as soon as the blastocyst is transferred to the equilibration medium.</li>
</ol>
5. Recording of the Blastocyst at the Equilibration Phase
<ol>
	<li>Transfer the microdroplet dish with the blastocyst to a camera-equipped microscope. Use the same magnification as previously for taking a snapshot of the same blastocyst. Position the microdroplet dish so that the blastocyst is positioned approximately in the center of the recording.</li>
	<li>Start recording with the microscope recording software. Note the countdown time at which the recording is started. 
	Note: See representative results of recordings of an intact (Figure 1, Video 1) and a collapsed blastocyst (Figure 2, Video 2) during the 10-min exposure to an equilibration solution.</li>
</ol>
6. Vitrification of the Blastocyst
<ol>
	<li>Aseptically dispense a 50-&#181;L drop of vitrification solution (M-199 HEPES-buffered medium with 15% DMSO, 15% ethylene glycol, 20% DSS, 0.5 M sucrose) in a Petri dish 2 min before the countdown ends.</li>
	<li>Stop recording when the 10-min countdown ends and quickly transfer the blastocyst to vitrification solution for 30 s at room temperature.</li>
	<li>Gently pipette the blastocyst at least 1x within the vitrification solution.</li>
	<li>Use a vitrification straw for loading the blastocyst. Load, seal, and plunge the vitrification straw into liquid nitrogen (LN) within 80 s, not exceeding 110 s after the initial exposure to the vitrification solution.</li>
</ol>
7. Video Editing of the Recorded Blastocyst During the Equilibration Phase
<ol>
	<li>Import a recorded video file into video editing software.</li>
	<li>Move the timeline slider to 20 s. Note the frame number at this time. 
	Note: This number will be used to decimate unnecessary video frames. Only frames every 20 s will be used.</li>
	<li>Click the Video tab, then Frame rate.... Under Frame rate conversion, check the Decimate by field and enter the previously noted frame number. Confirm by clicking OK. Click File &#8594; Export &#8594; Image sequence.</li>
	<li>Choose JPEG as the output format, set the directory to hold the saved sequence of images, and click OK. 
	Note: This will create a directory with up to 30 images of the recorded blastocyst in sequence during the equilibration phase of vitrification.</li>
</ol>
8. Measurements of the Blastocyst&#8217;s Cross-sectional Area from Images Created During the Equilibration Phase
<ol>
	<li>Open the video analysis software. Click the Window tab and select Timeline.</li>
	<li>In the Timeline pane, click on the film strip sign and choose Add Media... to add an image of the blastocyst before the equilibration phase of vitrification&#8212;and before the laser intervention, if there was any. 
	Note: This image will show the blastocyst at its most expanded state and its cross-sectional area will serve as a reference.</li>
	<li>Add the image sequence of the corresponding blastocyst generated previously with video editing software (of the equilibration phase of vitrification).</li>
	<li>Go to Image &#8594; Analysis &#8594; Select Data Points &#8594; Custom to open the Select data points window.</li>
	<li>Deselect all data points, then select only Area, and confirm by clicking OK.</li>
	<li>Move the timeline slider in the Timeline window to the first image.</li>
	<li>Click the Window tab and choose Tools. 
	Note: This will activate the Tools panel on the left side.</li>
	<li>Choose the Quick Selection Tool.</li>
	<li>Position the tool marker inside the blastocyst to touch the edge of the blastocyst. Outline the blastocyst by placing the tool marker on the blastocyst&#8217;s outer edge, clicking and holding the left mouse button, and dragging the tool marker along the blastocyst&#8217;s outer edge until the whole blastocyst&#8217;s cross-sectional area is selected. 
	Note: Zona pellucida is not a part of the measurement, so it must be excluded (Figure 5).</li>
	<li>In the Timeline window, click the Measurement Log and press the Record Measurements button. 
	Note: This will record the selected area.</li>
	<li>Return back to the Timeline, right-click on the image, and choose Deselect.</li>
	<li>Move the timeline slider to the next image and click on the corresponding tile beneath it.</li>
	<li>Outline the blastocyst in the new image with the Quick Selection Tool. Repeat the measurement.</li>
	<li>Repeat this procedure (steps 8.9 - 8.13) image by image until all the images&#8217; cross-sectional areas are measured and recorded.</li>
	<li>In the end, go to the Measurements Log, select all the measurements under the Area variable, and export them in a .txt file. 
	Note: The units of the cross-sectional area are square pixels.</li>
</ol>
9. Editing of the Data File with the Recorded Blastocyst&#8217;s Cross-sectional Areas from Images Created During the Equilibration Phase
<ol>
	<li>Transfer the data of the recorded blastocyst&#8217;s cross-sectional areas from the .txt file to a spreadsheet editor.</li>
	<li>Use the first cross-sectional area value as reference value 1 and express (transform) all the other values to be a fraction of that reference value.</li>
	<li>Create a new variable Area_r and paste the transcoded area values (except the reference value) under it.</li>
	<li>Next to the Area_r variable, create a Time variable and add the first time value. 
	Note: The first time value represents the time which passed from the moment the blastocyst was exposed to the equilibration solution to the onset of recording under a camera-equipped microscope.</li>
	<li>Calculate each next consecutive time value by adding 20 s to the previous time value. 
	Note: This is the time interval used at decimating unnecessary video frames created during the recording.</li>
</ol>
10. Warming of the Blastocyst
<ol>
	<li>Prepare media for warming.
	<ol>
		<li>A day before warming the blastocyst, aseptically dispense three 150-&#181;L drops of recovery medium in a 4-well dish. Also, aseptically dispense recovery medium in a microdroplet 9-well dish specially designed for time-lapse microscopy and recording. Cover the recovery medium with paraffin oil and preincubate it at 37 &#176;C with 6% CO2 and 5% O2. 
		Note: Recovery medium is a cultivation medium for blastocyst-stage embryos with added human serum albumin (HSA). The HSA in the recovery medium should be 12 mg/mL. To fill the holes in the 9-well dish, use a pipette for oocyte denudation to avoid the creation of air bubbles, then overlay the microdroplet area with 30 &#181;L of recovery medium.</li>
		<li>On the day of warming the blastocyst, aseptically dispense 500 &#181;L of thawing solution (TS) in a single well of a 4-well dish and allow it to warm to 37 &#176;C in an incubator without CO2.</li>
		<li>At room temperature, aseptically dispense one 50-&#181;L drop of dilution solution (DS) and two 50-&#181;L drops of washing solution (WS) in a sterile Petri dish. Cover the DS and the WS1 and WS2 drops with paraffin oil. 
		Note: Instead of a Petri dish, a 4-well dish can also be used.</li>
	</ol>
	</li>
	<li>Warm the blastocyst.
	<ol>
		<li>Identify the HSV straw with the vitrified blastocyst to be removed from LN storage and quickly transfer the straw to an LN-filled portable reservoir in preparation for the warming procedure.</li>
		<li>Lift the straw with forceps, just enough to expose the colored handling rod. Use a self-adjusting wire stripper to cut the straw at the height of the colored handling rod.</li>
		<li>Grab the handling rod and extract it from the straw with a swift, yet controlled movement, and immediately plunge the curved spatula of the handling rod into the 37 &#176;C TS.</li>
		<li>Gently swirl the handling rod to detach the blastocyst and leave it for a total of 1 min in TS.</li>
		<li>At room temperature, transfer the blastocyst consecutively to DS, WS1, and WS2 for 4 min in each medium.</li>
		<li>After the warming procedure, transfer the blastocyst to a 4-well dish with recovery medium and wash it in all three drops by gently pipetting it. 
		Note: The washing is carried out in approximately 1 min.</li>
		<li>Transfer the blastocyst to the time-lapse 9-well dish with recovery medium. Place the 9-well dish under a time-lapse camera in an incubator (at 37 &#176;C with 6% CO2 and 5% O2). 
		Note: Thre warming procedure, which continues with placing the 9-well dish under a time-lapse camera in an incubator, lasts approximately 17 min.</li>
	</ol>
	</li>
</ol>
11. Time-lapse Recording of the Blastocyst Re-expansion During Recovery Post-warming
<ol>
	<li>Run time-lapse recording software.</li>
	<li>Choose a recording camera.</li>
	<li>Press the Live mode button.</li>
	<li>Place the mouse cursor on the image and use the scroll button to enlarge it. Click and hold the left mouse button and move the cursor to place the well with the blastocyst in the middle of the screen.</li>
	<li>Under Focusing, use the up-down green arrows to focus the recording plane of the blastocyst. Under Light intensity, set the light intensity.</li>
	<li>Click the Microscope parameters button to set the Exposure time and Gamma.</li>
	<li>Press Close live mode.</li>
	<li>Press the Start project button and enter the project data, select the culture dish type (3 x 3 or 4 x 4) and uncheck all positions except the one to be recorded.</li>
	<li>Set the capture timing to take a picture every 5 min.</li>
	<li>Start recording by pressing the Approve button. Record at least 150 min.</li>
	<li>Stop recording by pressing the Stop project button. 
	Note: See the representative results of the recording of an intact (Figure 3) and a collapsed blastocyst (Figure 4) during the post-warming recovery period.</li>
</ol>
12. Video Editing of the Recorded Blastocyst During the Re-expansion Post-warming
<ol>
	<li>Go to the project folder and locate recorded images that are roughly 80 KB in size.</li>
	<li>Create another folder and transfer these images into it. Rename the images in numbers according to the time created. Import them into the video editing software as an image sequence.</li>
	<li>Go to Video tab &#8594; Filter &#8594; Add and select Null transform filter.</li>
	<li>Click the Cropping button on the right side and crop the image, leaving visible only the field with the recorded blastocyst. Confirm with OK and again with OK.</li>
	<li>Click File &#8594; Export &#8594; Image sequence. Choose JPEG as the output format, set the directory to hold the saved sequence of images, and click OK. 
	Note: This will create resized (cropped) images prepared for further measurements in the video analysis software.</li>
</ol>
13. Setting of a Measurement Scale in Video Analysis Software for the Images Created with the Time-lapse Recording Software
<ol>
	<li>Run the Analyzer of the time-lapse recording software.</li>
	<li>Open the project with the recorded blastocyst.</li>
	<li>Click the Analyze button on the field with the recorded blastocyst. 
	Note: A new Analyze window will open. Click on the Measurement button to show the measurement tool.</li>
	<li>Use the measurement tool to measure the width of the whole image.</li>
	<li>Right-click on the image and save it. In the properties, check the image&#8217;s width in pixels.</li>
	<li>Divide the image&#8217;s actual width with its width in pixels. 
	Note: This calculates the actual length of a single pixel in this image.</li>
	<li>Run the video analysis software.</li>
	<li>Click the Window tab and select Timeline.</li>
	<li>In the Timeline pane, click on the film strip sign and choose Add Media... to add the image sequence of the post-warm re-expanding blastocyst.</li>
	<li>Click Image &#8594; Analysis &#8594; Set Measurement Scale &#8594; Custom to open the Measurement Scale window. For Pixel length, enter 1; for Logical length, enter the previously calculated actual length of a pixel (step 13.6); for Logical units, enter &#181;m. Save the preset.</li>
</ol>
14. Measurements of the Blastocyst&#8217;s Cross-sectional Area from Images Created During the Re-expansion Post-warm
<ol>
	<li>Once the Measurement scale is set and the image sequence imported, measure the blastocyst&#8217;s cross-sectional areas the same way as described in step 8, with the only difference being that the measured cross-sectional areas in these measurements have units in &#181;m2.</li>
</ol>
15. Editing of the Data File with the Recorded Blastocyst&#8217;s Cross-sectional Areas from Images Created during the Re-expansion Post-warm
<ol>
	<li>Transfer the data of the recorded blastocyst&#8217;s cross-sectional areas from the .txt file to a spreadsheet editor.</li>
	<li>Create a Time variable next to the Area variable and add the first time value. 
	Note: In this case, the first time value is set to 0 minutes, which is the onset of the time-lapse recording. Each next consecutive time value is calculated by adding 5 minutes to the previous time value. Five minutes is the time interval used between two consecutive time-lapse recordings.</li>
</ol>
16. Creation of a Line Diagram
<ol>
	<li>Import the spreadsheet data file in statistical analysis software for creating a time plot of the blastocyst&#8217;s cross-sectional area changes in time during the equilibration phase of the vitrification or re-expansion post-warming.</li>
</ol>

<ol>
	<li>Trounson, A., Mohr, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+pregnancy+following+cryopreservation,+thawing+and+transfer+of+an+eight-cell+embryo.">Human pregnancy following cryopreservation, thawing and transfer of an eight-cell embryo.</a> Nature. 305 (5936), 707-709 (1983).</li><li>Leibo, S. P., McGrath, J. J., Cravalho, E. 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A., Choi, A., Orief, Y., Agarwal, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factors+affecting+the+outcome+of+human+blastocyst+vitrification.">Factors affecting the outcome of human blastocyst vitrification.</a> Reproductive Biology and Endocrinology. 16 (7), 99 (2009).</li><li>Ebner, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphokinetics+of+vitrified+and+warmed+blastocysts+predicts+implantation+potential.">Morphokinetics of vitrified and warmed blastocysts predicts implantation potential.</a> Journal of Assisted Reproduction and Genetics. 34 (2), 239-244 (2017).</li><li>Coello, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+morphological+dynamics+of+blastocysts+after+vitrification/warming:+defining+new+predictive+variables+of+implantation.">Analysis of the morphological dynamics of blastocysts after vitrification/warming: defining new predictive variables of implantation.</a> Fertility Sterility. 108 (4), 659-666 (2017).</li><li>Huang, T. T. F., Chinn, K., Kosasa, T., Ahn, H. J., Kessel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphokinetics+of+human+blastocyst+expansion+in+vitro.">Morphokinetics of human blastocyst expansion in vitro.</a> Reproductive Biomedicine Online. 33, 659-667 (2016).</li><li>Vanderzwalmen, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lower+intracellular+concentration+of+cryoprotectants+after+vitrification+than+after+slow+freezing+despite+exposure+to+higher+concentration+of+cryoprotectant+solutions.">Lower intracellular concentration of cryoprotectants after vitrification than after slow freezing despite exposure to higher concentration of cryoprotectant solutions.</a> Human Reproduction. 28, 2101-2110 (2013).</li></ol>

The authors have nothing to disclose.

The protocol for the observation of blastocyst morphodynamics during and after cryopreservation can also be carried out by using similar instruments and software tools from other manufacturers. Time-lapse systems adjusted for embryology allow the continuous monitoring of embryo development. The purpose of this work was to introduce the quantification of blastocyst behavior during the preparation of blastocysts for vitrification and after their warming. This was done by the objective measurement of changes in the morpholo...

Cryopreservation of human preimplantation embryos from the in vitro fertilization program (IVF) is nowadays a routine practice in most IVF laboratories. The slow embryo freezing method began to be clinically used in 1985, with the introduction of specific cryoprotectants and computer-controlled freezers, which enabled the controlled cooling of embryos down to -7 &#176;C, when ice nucleation (seeding) was induced in the surrounding cryoprotective medium1. By continuous cooling, the ice crystals would grow, causing the hyperosmolality of the remaining liquid fraction and, consequently, the dehydration and shrinkage of the embryonic cells. At -30 &#176;C or -80 &#176;C, the embryos would be plunged into the liquid nitrogen for longer storage. What was happening with the embryos or oocytes during cooling could be observed only by specially adapted cryomicroscopes, which helped to improve cryopreservation protocols2. The freezing of blastocysts, a higher-volume and more fluid-contained embryonic stage, gave less promising clinical results in those days3.
The breakthrough in the cryopreservation of the blastocyst was the introduction of the vitrification method, in which the dehydration of the cells took place before cooling by using high concentrations of cryoprotectants4. The removal of the fluid from the blastocoel before vitrification can also be achieved by making a mechanical opening between two trophectoderm cells5. Although the immediate blastocyst survival rate after vitrification and warming is above 90%, and the clinical outcome following the transfer of vitrified/warmed blastocyst into the uterine cavity is almost comparable to results after the transfer of fresh embryos, this cryopreservation method has not yet been standardized6,7. Vitrification protocols vary according to (a) the type and concentration of the cryoprotectants, (b) the number of previtrification steps, (c) the duration of individual steps, (d) the use of an artificial blastocoel collapsing before vitrification or not, (e) collapsing methods, (f) the blastocyst expansion stage, and (g) the equilibration/vitrification temperature at which the embryos should be vitrified8. Since cryoprotectants can be toxic for cells, the blastocyst exposure time to these solutions has to be well defined. However, some cryopreservation media manufacturers allow very flexible protocols.
The interest of scientists is usually focused on studying the blastocyst re-expansion ability with the aim to find new biomarkers with a better prediction of implantation6,9,10. How human blastocysts dehydrate at different steps of adding cryoprotectants before vitrification and what happens with blastocysts after vitrification and warming, when cryoprotectants have to be removed from the cells, and how the blastocysts rehydrate and re-expand after warming, is not well described nor understood. The development of a methodology for the objective and quantified monitoring of blastocyst behavior during different cryopreservation steps is, thus, rationale.
With time-lapse microscopes of various manufacturers, it is now possible to monitor the behavior of the blastocyst in pre- and post-vitrification phases. By including additional computer tools, measurements of their size (morphometry) can also be performed. By measuring the decrease or increase in embryo size at a given time, it is possible to objectify the evaluation of morphodynamics of embryos during dehydration and rehydration.

<table>
	<tbody>
		<tr>
			<td>Inverted microscope Eclipse TE2000-U&#160;</td>
			<td>Nikon, Japan</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Saturn 5 Laser System</td>
			<td>Research Instruments, Origio, Denmark</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital camera DC1</td>
			<td>Research Instruments, Origio, Denmark</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital camera DC2</td>
			<td>Research Instruments, Origio, Denmark</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Cronus 3.7</td>
			<td>Research Instruments, Origio, Denmark</td>
			<td>/</td>
			<td>microscope recording software</td>
		</tr>
		<tr>
			<td>Incubator with 6% CO2, 5% O2</td>
			<td>Binder, Germany</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Primo Vision microscope</td>
			<td>Vitrolife, Sweden</td>
			<td>16600</td>
			<td></td>
		</tr>
		<tr>
			<td>Primo Vision Capture software</td>
			<td>Vitrolife, Sweden</td>
			<td>16608</td>
			<td>time-lapse recording software</td>
		</tr>
		<tr>
			<td>Adobe Photoshop CS6 Extended software</td>
			<td>Adobe Systems Incorporated, USA</td>
			<td>/</td>
			<td>video analysis software</td>
		</tr>
		<tr>
			<td>VirtualDub&#160;</td>
			<td>Avery Lee</td>
			<td>/</td>
			<td>video editing software</td>
		</tr>
		<tr>
			<td>Microsoft Office Excell&#160;</td>
			<td>Microsoft, USA</td>
			<td>/</td>
			<td>spreadsheet editor</td>
		</tr>
		<tr>
			<td>PrimoVision culture dish</td>
			<td>Vitrolife, Sweden</td>
			<td>16604</td>
			<td></td>
		</tr>
		<tr>
			<td>G2-plus medium</td>
			<td>Vitrolife, Sweden</td>
			<td>10132</td>
			<td>cultivation medium for blastocyst stage embryos</td>
		</tr>
		<tr>
			<td>Human Serum Albumins</td>
			<td>Vitrolife, Sweden</td>
			<td>10064</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin oil</td>
			<td>Vitrolife, Sweden</td>
			<td>10029</td>
			<td></td>
		</tr>
		<tr>
			<td>Equilibration solution medium</td>
			<td>Irvine Scientific, Ireland</td>
			<td>90131</td>
			<td></td>
		</tr>
		<tr>
			<td>Vitrification solution medium</td>
			<td>Irvine Scientific, Ireland</td>
			<td>90132</td>
			<td></td>
		</tr>
		<tr>
			<td>Thawing solution medium</td>
			<td>Irvine Scientific, Ireland</td>
			<td>90134</td>
			<td></td>
		</tr>
		<tr>
			<td>Dilution solution medium</td>
			<td>Irvine Scientific, Ireland</td>
			<td>90135</td>
			<td></td>
		</tr>
		<tr>
			<td>Washing solution medium</td>
			<td>Irvine Scientific, Ireland</td>
			<td>90136</td>
			<td></td>
		</tr>
		<tr>
			<td>HSV Vitrification straws</td>
			<td>CryoBio System, France</td>
			<td>025246, 025249, 025250, 025248</td>
			<td></td>
		</tr>
		<tr>
			<td>Liquid nitrogen</td>
			<td>/</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cryo vessel Biosafe 120 MD &#946;</td>
			<td>Cryotherm, Germany</td>
			<td>229286</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryo tank</td>
			<td>Cryotherm, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>/</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Scisors</td>
			<td>/</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Pippete for blastocyst manipulation</td>
			<td>Gynetics, Belgium</td>
			<td>ID275/10</td>
			<td>diameter 275 &#181;m</td>
		</tr>
		<tr>
			<td>Pipette for oocyte denudation</td>
			<td>Vitromed, Germany</td>
			<td>V-DEN-135</td>
			<td>diameter 135 &#181;m</td>
		</tr>
		<tr>
			<td>Pipettor EZ-Grip</td>
			<td>Research Instruments</td>
			<td>7-72-2802</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital interval timer Assistent</td>
			<td>Glaswarenfabrik Karl Hecht</td>
			<td>41977010</td>
			<td></td>
		</tr>
		<tr>
			<td>IBM SPSS Statistics 21</td>
			<td>IBM, USA</td>
			<td>/</td>
			<td>statistical analysis software</td>
		</tr>
		<tr>
			<td>Self adjusting wire stripper</td>
			<td>Knipex, Germany</td>
			<td>1262180</td>
			<td></td>
		</tr>
	</tbody>
</table>

morphometric protocol for the objective assessment of blastocyst behavior during vitrification and warming steps

This article describes the noninvasive method of blastocyst morphometry based on time-lapse microphotography for the accurate monitoring of a blastocyst&#39;s volume changing during individual phases before and after vitrification. The method can be useful in searching for the most optimal timing of blastocyst exposure to different concentrations of cryoprotectants by observing blastocyst shrinkage and re-expansion in different pre- and post-vitrification phases. With this methodology, the blastocyst vitrification protocol can be optimized. For a better demonstration of the usefulness of this morphometric method, two different blastocyst preparation protocols for vitrification are compared; one with using an artificial blastocoel collapsing and one without this intervention before vitrification. Both blastocysts&#39; volume changes are followed by time-lapse microphotography and measured by photo-editing software tools. The measurements are taken every 20 seconds in previtrification phases and every 5 minutes in the post-warming period. The changes of the blastocyst dimensions per time unit are presented graphically in line diagrams. The results show a long equilibration previtrification phase in which the intact blastocyst first shrinks and then slowly refills the blastocoel, entering vitrification with a fluid-filled blastocoel. The artificially collapsed blastocyst remains in its shrunken stage through the entire equilibration phase. During the vitrification phase, it also does not change its volume. Since the blastocyst morphometry shows a constant volume of the artificially collapsed blastocysts during the previtrification step, it seems that this stage could be shorter. The described protocol provides many additional comparative parameters of blastocyst behavior during and after cryopreservation on the basis of the speed and intensity of the volume changes, the number of partial blastocoel contractions or total blastocyst collapses, and the time to a total blastocoel re-expansion or the time to hatching.

In a demonstration, we showed blastocyst morphodynamics in only one previtrification and one post-warming phase. A difference in the blastocyst&#39;s volume at the end of the equilibration phase and at the beginning of recovering in culture medium showed the intensity of embryo shrinkage, which is, in fact, the intensity of embryo preservation against ice crystallization.
As it can be noticed from Figure 1</...

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Morphometric Protocol for the Objective Assessment of Blastocyst Behavior During Vitrification and Warming Steps

Here we present a time-lapse morphometric protocol to follow the intensity of blastocyst shrinkage and re-expansion during previtrification interventions and post-warming recovery. The application of the protocol is possible in in vitro fertilization laboratories equipped with time-lapse microscopes and is recommended in the development of an optimal blastocyst vitrification method.

This article describes the noninvasive method of blastocyst morphometry based on time-lapse microphotography for the accurate monitoring of a ...

morphometric-protocol-for-objective-assessment-blastocyst-behavior

Research

JoVE Journal

Bioengineering

8.6K Views.  University Medical Centre Maribor. Here we present a time-lapse morphometric protocol to follow the intensity of blastocyst shrinkage and re-expansion during previtrification interventions and post-warming recovery. The application of the protocol is possible in in vitro fertilization laboratories equipped with time-lapse microscopes and is recommended in the development of an optimal blastocyst vitrification method.

Video: Morphometric Protocol for the Objective Assessment of Blastocyst Behavior During Vitrification and Warming Steps

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the arterial wall lead to tissue injury, which initiates restenosis2-7. This hypothesis needs further investigations including better quantifications of non-uniform strain distribution on the artery following stent implantation. A non-contact surface strain measurement method for the stented artery is presented in this work. ARAMIS stereo optical surface strain measurement system uses two optical high speed cameras to capture the motion of each reference point, and resolve three dimensional strains over the deforming surface8,9. As a mesh stent is deployed into a latex vessel with a random contrasting pattern sprayed or drawn on its outer surface, the surface strain is recorded at every instant of the deformation. The calculated strain distributions can then be used to understand the local lesion response, validate the computational models, and formulate hypotheses for further in vivo study.

Stent-induced arterial strain distributions are characterized using an optical surface strain measurement system. This visualization technique is used to gain insights into the impact of stent implantation on the host vessel.

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the ...

Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of tri-leaflet polymer valve prostheses. However, identification of a protocol for initial assessment of polymer valve hydrodynamic functionality is paramount during the early stages of the design process. Traditional in vitro pulse duplicator systems are not configured to accommodate flexible tri-leaflet materials; in addition, assessment of polymer valve functionality needs to be made in a relative context to native and prosthetic heart valves under identical test conditions so that variability in measurements from different instruments can be avoided. Accordingly, we conducted hydrodynamic assessment of i) native (n = 4, mean diameter, D = 20 mm), ii) bi-leaflet mechanical (n= 2, D = 23 mm) and iii) polymer valves (n = 5, D = 22 mm) via the use of a commercially available pulse duplicator system (ViVitro Labs Inc, Victoria, BC) that was modified to accommodate tri-leaflet valve geometries. Tri-leaflet silicone valves developed at the University of Florida comprised the polymer valve group. A mixture in the ratio of 35:65 glycerin to water was used to mimic blood physical properties. Instantaneous flow rate was measured at the interface of the left ventricle and aortic units while pressure was recorded at the ventricular and aortic positions. Bi-leaflet and native valve data from the literature was used to validate flow and pressure readings. The following hydrodynamic metrics were reported: forward flow pressure drop, aortic root mean square forward flow rate, aortic closing, leakage&#160;and regurgitant volume, transaortic closing, leakage, and total energy losses. Representative results indicated that hydrodynamic metrics from the three valve groups could be successfully obtained by incorporating a custom-built assembly into a commercially available pulse duplicator system and subsequently, objectively compared to provide insights on functional aspects of polymer valve design.

There has been renewed interest in developing polymer valves. Here, the objectives are to demonstrate the feasibility of modifying a commercial pulse duplicator to accommodate tri-leaflet geometries and to define a protocol to present polymer valve hydrodynamic data in comparison to native and prosthetic valve data collected under near-identical conditions.

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of ...

Engineering Platform and Experimental Protocol for Design and Evaluation of a Neurally-controlled Powered Transfemoral Prosthesis

To enable intuitive operation of powered artificial legs, an interface between user and prosthesis that can recognize the user's movement intent is desired. A novel neural-machine interface (NMI) based on neuromuscular-mechanical fusion developed in our previous study has demonstrated a great potential to accurately identify the intended movement of transfemoral amputees. However, this interface has not yet been integrated with a powered prosthetic leg for true neural control. This study aimed to report (1) a flexible platform to implement and optimize neural control of powered lower limb prosthesis and (2) an experimental setup and protocol to evaluate neural prosthesis control on patients with lower limb amputations. First a platform based on a PC and a visual programming environment were developed to implement the prosthesis control algorithms, including NMI training algorithm, NMI online testing algorithm, and intrinsic control algorithm. To demonstrate the function of this platform, in this study the NMI based on neuromuscular-mechanical fusion was hierarchically integrated with intrinsic control of a prototypical transfemoral prosthesis. One patient with a unilateral transfemoral amputation was recruited to evaluate our implemented neural controller when performing activities, such as standing, level-ground walking, ramp ascent, and ramp descent continuously in the laboratory. A novel experimental setup and protocol were developed in order to test the new prosthesis control safely and efficiently. The presented proof-of-concept platform and experimental setup and protocol could aid the future development and application of neurally-controlled powered artificial legs.

Neural-machine interfaces (NMI) have been developed to identify the user's locomotion mode. These NMIs are potentially useful for neural control of powered artificial legs, but have not been fully demonstrated. This paper presented (1) our designed engineering platform for easy implementation and development of neural control for powered lower limb prostheses and (2) an experimental setup and protocol in a laboratory environment to evaluate neurally-controlled artificial legs on patients with lower limb amputations safely and efficiently.

To enable intuitive operation of powered artificial legs, an interface between user and prosthesis that can recognize the user's movement intent is ...

Expanding Nanopatterned Substrates Using Stitch Technique for Nanotopographical Modulation of Cell Behavior

Substrate nanotopography has been shown to be a potent modulator of cell phenotype and function. To dissect nanotopography modulation of cell behavior, a large area of nanopatterned substrate is desirable so that enough cells can be cultured on the nanotopography for subsequent biochemical and molecular biology analyses. However, current nanofabrication techniques have limitations to generate highly defined nanopatterns over a large area. Herein, we present a method to expand nanopatterned substrates from a small, highly defined nanopattern to a large area using stitch technique. The method combines multiple techniques, involving soft lithography to replicate poly(dimethylsiloxane) (PDMS) molds from a well-defined mold, stitch technique to assemble multiple PDMS molds to a single large mold, and nanoimprinting to generate a master mold on polystyrene (PS) substrates. With the PS master mold, we produce PDMS working substrates and demonstrate nanotopographical modulation of cell spreading. This method provides a simple, affordable yet versatile avenue to generate well-defined nanopatterns over large areas, and is potentially extended to create micro-/nanoscale devices with hybrid components.

A protocol for producing a large area of nanopatterned substrate from small nanopatterned molds for study of nanotopographical modulation of cell behavior is presented.

Substrate nanotopography has been shown to be a potent modulator of cell phenotype and function. To dissect nanotopography modulation of cell behavior, a ...

Dissection, MicroCT Scanning and Morphometric Analyses of the Baculum

Modern morphometrics provides powerful methods to quantify size and shape variation. A basic requirement is a list of coordinates that define landmarks; however such coordinates must represent homologous structures across specimens. While many biological objects consist of easily identified landmarks to satisfy the assumption of homology, many lack such structures. One potential solution is to mathematically place semi-landmarks on an object that represent the same morphological region across specimens. Here, we illustrate a recently developed pipeline to mathematically define semi-landmarks from the mouse baculum (penis bone). Our methods should be applicable to a wide range of objects.

Many biological structures lack easily definable landmarks, making it difficult to apply modern morphometric methods. Here we illustrate methods to study the mouse baculum (a bone in the penis), including dissection and microCT scanning, followed by computational methods to define semi-landmarks that are used to quantify size and shape variation.

Modern morphometrics provides powerful methods to quantify size and shape variation. A basic requirement is a list of coordinates that define landmarks; ...

Protocol for MicroRNA Transfer into Adult Bone Marrow-derived Hematopoietic Stem Cells to Enable Cell Engineering Combined with Magnetic Targeting

While CD133+ hematopoietic stem cells (SCs) have been proven to provide high potential in the field of regenerative medicine, their low retention rates after injection into injured tissues as well as the observed massive cell death rates lead to very restricted therapeutic effects. To overcome these limitations, we sought to establish a non-viral based protocol for suitable cell engineering prior to their administration. The modification of human CD133+ expressing SCs using microRNA (miR) loaded magnetic polyplexes was addressed with respect to uptake efficiency and safety as well as the targeting potential of the cells. Relying on our protocol, we can achieve high miR uptake rates of 80&#8211;90% while the CD133+ stem cell properties remain unaffected. Moreover, these modified cells offer the option of magnetic targeting. We describe here a safe and highly efficient procedure for the modification of CD133+ SCs. We expect this approach to provide a standard technology for optimization of therapeutic stem cell effects and for monitoring of the administered cell product via magnetic resonance imaging (MRI).

This protocol illustrates a safe and efficient procedure to modify CD133+ hematopoietic stem cells. The presented non-viral, magnetic polyplex-based approach may provide a basis for the optimization of therapeutic stem cell effects as well as for monitoring the administered cell product via magnetic resonance imaging.

While CD133+ hematopoietic stem cells (SCs) have been proven to provide high potential in the field of regenerative medicine, their low retention rates ...

Environmental Dynamic Mechanical Analysis to Predict the Softening Behavior of Neural Implants

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior of these materials. In the past, it has not been possible to satisfactorily measure the softening of thin films under conditions mimicking body environment without substantial effort. This publication presents a new and simple method that allows dynamic mechanical analysis (DMA) of polymers in solutions, such as phosphate buffered saline (PBS), at relevant temperatures. The use of environmental DMA allows measurement of the softening effects of polymers due to plasticization in various media and temperatures, which therefore allows a prediction of the materials behavior under in vivo conditions.

To allow reliable predictions of the softening of polymeric substrates for neural implants in an in vivo environment, it is important to have a reliable in vitro method. Here, the use of dynamic mechanical analysis in phosphate buffered saline at body temperature is presented.

When using dynamically softening substrates for neural implants, it is important to have a reliable in vitro method to characterize the softening behavior ...

Bulk Droplet Vitrification for Primary Hepatocyte Preservation

Vitrification is a promising ice-free alternative for classic slow-freezing (at approximately 1 &#176;C/min) cryopreservation of biological samples. Vitrification requires extremely fast cooling rates to achieve transition of water into the glass phase while avoiding injurious ice formation. Although pre-incubation with cryoprotective agents (CPA) can reduce the critical cooling rate of biological samples, high concentrations are generally needed to enable ice-free cryopreservation by vitrification. As a result, vitrification is hampered by CPA toxicity and restricted to small samples that can be cooled fast. It was recently demonstrated that these inherent limitations can be overcome by bulk droplet vitrification. Using this novel method, cells are first pre-incubated with a low intracellular CPA concentration. Leveraging rapid osmotic dehydration, the intracellular CPA is concentrated directly ahead of vitrification, without the need to fully equilibrate toxic CPA concentrations. The cellular dehydration is performed in a fluidic device and integrated with continuous high throughput generation of large sized droplets that are vitrified in liquid nitrogen. This ice-free cryopreservation method with minimal CPA toxicity is suitable for large cell quantities and results in increased hepatocyte viability and metabolic function as compared to classical slow-freezing cryopreservation. This manuscript describes the methods for successful bulk droplet vitrification in detail.

This manuscript describes an ice-free cryopreservation method for large quantities of rat hepatocytes whereby primary cells are pre-incubated with cryoprotective agents at a low concentration and vitrified in large droplets.

Vitrification is a promising ice-free alternative for classic slow-freezing (at approximately 1 &#176;C/min) cryopreservation of biological samples. ...

Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the microscale&#8213;to create micro-environments for microbial chemotaxis experiments. To create chemical pulses, we developed a system to introduce amino acid sources near-instantaneously by photolysis of caged amino acids within a polydimethylsiloxane (PDMS) microfluidic chamber containing a bacterial suspension. We applied this method to the chemotactic bacterium, Vibrio ordalii, which can actively climb these dynamic chemical gradients while being tracked by video microscopy. Amino acids, rendered biologically inert (&#39;caged&#39;) by chemical modification with a photoremovable protecting group, are uniformly present in the suspension but not available for consumption until their sudden release, which occurs at user-defined points in time and space by means of a near-UV-A focused LED beam. The number of molecules released in the pulse can be determined by a calibration relationship between exposure time and uncaging fraction, where the absorption spectrum after photolysis is characterized by using UV-Vis spectroscopy. A nanoporous polycarbonate (PCTE) membrane can be integrated into the microfluidic device to allow the continuous removal by flow of the uncaged compounds and the spent media. A strong, irreversible bond between the PCTE membrane and the PDMS microfluidic structure is achieved by coating the membrane with a solution of 3-aminopropyltriethoxysilane (APTES) followed by plasma activation of the surfaces to be bonded. A computer-controlled system can generate user-defined sequences of pulses at different locations and with different intensities, so as to create resource landscapes with prescribed spatial and temporal variability. In each chemical landscape, the dynamics of bacterial movement at the individual scale and their accumulation at the population level can be obtained, thereby allowing the quantification of chemotactic performance and its effects on bacterial aggregations in ecologically relevant environments.

A protocol for the generation of dynamic chemical landscapes by photolysis within microfluidic and millifluidic setups is presented. This methodology is suitable to study diverse biological processes, including the motile behavior, nutrient uptake, or adaptation to chemicals of microorganisms, both at the single cell and population level.

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the ...

Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between ...