The authors acknowledge funding support from the National Science Foundation (NSF) STTR Phase 1 Award (#: 1938939), by ASME Defense Advanced Research Projects Agency (DARPA) Award (#: N660012024075), and Johns Hopkins Institute for Clinical and Translational Research&#39;s (ICTR&#39;s) Clinical Research Scholars Program (KL2), administered by the National Institutes of Health (NIH) National Center for Advancing Translational Sciences (NCATS). The cells were purchased from and provided by the Mayo Foundation for Medical Education and Research.

All animal studies were approved and conducted in accordance with the Johns Hopkins University Institutional Animal Care and Use Committee (IACUC). Athymic nude female mice (age: 10 weeks) were obtained from commercial sources (see Table of Materials). All Biosafety Level 2 (BSL-2) regulations were followed, including the usage of masks, gloves, and gowns.
1. Tumor implantations and bioluminescence imaging
<ol>
	<li>During the initial phase of the study, perform intracranial tumor implantation, following a previously published report6. 
	NOTE: In this study, 100,000 M59 human GBM xenograft cells in 4 &#181;L of cell suspension were used for implantation at a 3.0 mm depth into the skull.</li>
	<li>Prior to treatment, quantify the tumor size in each mouse noninvasively using an in vivo luminescence imaging system (see Table of Materials) following a previously published report7. 
	NOTE: This was done on the date of treatment, 7 days following initial tumor implantations.</li>
	<li>Using imaging quantification, divide the mice into comparable subgroups for treatment. 
	&#8203;NOTE: In this study, two subgroups were included: (1) untreated tumor-bearing mice (n = 4) and (2) tumor-bearing mice undergoing SDT (n = 4). No statistically significant difference in pre-treatment tumor size was observed between the two groups (p &#62; 0.05).</li>
</ol>
2. Treatment day setup
<ol>
	<li>Remove 5-ALA hydrochloride (see Table of Materials) from the freezer and weigh out 200 mg/kg mouse weight of 5-ALA (e.g., for a 25 g mouse, weigh out 5 mg.). Do so under ambient light only. 
	NOTE: Sterile all the equipment before use.&#160;</li>
	<li>Dissolve the total amount of 5-ALA in phosphate-buffered saline (PBS), such that each animal is administered a 60 mg/mL solution of 5-ALA with the correct weight dosage (e.g., for a 25 g mouse, dissolve 5 mg of 5-ALA into 83.33 &#181;L of PBS).</li>
	<li>For each animal in the SDT testing group, inject the proper amount of solution of 5-ALA into the animal intraperitoneally (calculated in steps 2.1 and 2.2).</li>
	<li>Leave the animals in their cages for 3 h to metabolize the 5-ALA.</li>
</ol>
3. Preparation of requisites for experiments
<ol>
	<li>Fill a reservoir with deionized (DI) water. 
	NOTE: The amount of water needed is based on the number of mice used in the study.</li>
	<li>Attach the Flow in and Flow out tubes to the ultrasonic degasser (see Table of Materials) and plug in the degasser to the power (1 = ON, 0 = OFF).</li>
	<li>Place the other end of the Flow in and Flow out tubes into the DI water reservoir.</li>
	<li>Turn on the degasser and run for 45 min.</li>
	<li>Fill the newly degassed water to the top of an airtight, sealed container to be used during the study.</li>
	<li>Pour ultrasound gel (see Table of Materials) into a conical tube, and then place into a centrifuge and spin at 160 x g for 5 min to remove any air from the gel. 
	&#8203;NOTE: Enough gel is needed to form a dollop of gel on the head of each animal.</li>
</ol>
4. MRgFUS system setup
<ol>
	<li>Connect the MRgFUS system (see Table of Materials) using the wiring diagram as seen in Figure 1 (bottom panel).
	<ol>
		<li>Connect all necessary components (desktop computer, monitor, MRgFUS platform, oscilloscope, amplifier) to power.</li>
		<li>Connect all cables to the correct locations.</li>
		<li>Connect the desired transducer to the MRgFUS platform via the BNC and coaxial cables, and then connect the corresponding impedance matching box to the correct wires. 
		NOTE: A 515 kHz transducer was used for the present study.</li>
	</ol>
	</li>
	<li>Turn on all devices.</li>
	<li>On the desktop computer operating system, open the AUREUS application, which is already integrated into the FUS system software.</li>
	<li>Select Connect All Hardware to connect the hardware to the system and make sure that the components are communicating with the software.</li>
	<li>Select the transducer used by selecting the drop-down menu and choosing the desired transducer.</li>
</ol>
5. Initialization
<ol>
	<li>Unscrew the bottom screw from the phantom and fill the cavity with deionized and degassed water until it overflows, and then replace the screw again.</li>
	<li>Insert the phantom in its corresponding MRI bed location (see Figure 2), and then place the MRI bed into the MRI cradle in its corresponding slot.</li>
	<li>Place the MRI cradle in its corresponding location in the MRI scanner (see Table of Materials). Adjust the cradle so that the phantom MRI bed can slide into the MRI bore without any obstructions to obtain a high-quality image of the phantom. Mark this location so that it is easily replicable in the future. 
	NOTE: Once this location is found, the location will not be changed for the remainder of the treatment. Therefore, make sure that it is an adequate location for animal MRI placement, or the entire registration will need to be repeated.</li>
	<li>Take an MRI scan of the phantom using the settings in Table 1.</li>
	<li>Remove the cradle from the magnet bore but keep it on the scanner. Remove the MRI bed containing the phantom from the cradle, and then place the bed with the phantom onto the MRgFUS system by sliding the peg on the bottom into its correct slot.</li>
	<li>Slot the registration tip onto the magnetic slots on the transducer arm so that it points downward toward the phantom (see Figure 2).</li>
	<li>On the software, choose Select a New Home Position, and then select Jog Mode On to begin guided focused finding. After jog mode is toggled, use the left, right, up, down, page up, and page down keys to manually move the transducer arm around in the left, right, forward, backward, up, and down directions, respectively.</li>
	<li>Adjust the pointer manually in all three dimensions until the tip of the pointer touches the middle of the cross pattern that lies on the top of the phantom (see Figure 2).</li>
	<li>Download the phantom MRI images to the computer and place them in a folder in the chosen directory, and then load the MRI images into the software by selecting Load Phantom Images. The axial phantom slices will then populate the screen on the right side. Scroll through these slices via the mouse scroll bar. Adjust the brightness by clicking and holding on the image and then moving the mouse up or down. 
	NOTE: If any of the files in the folder are not uncompressed DICOM files, then the software will not be able to read and import them, so remove any other files from that folder.</li>
	<li>Scroll to any image of the phantom where there is a clear circle with three dark holes in each. Click on the middle of the phantom, and a red circle will pop up. Click and drag on the circle until it is the same diameter and lines up with the circumference of the phantom (see Figure 2).</li>
	<li>The coordinates of this position are referred to as &#39;home position&#39;; save this by clicking Set L/R, A/P, and S/I.</li>
	<li>Click Jog Mode Off, remove the registration tip from the transducer arm, and then select Exit Focus Finding. Confirm the home position to complete the initialization sequence.</li>
</ol>
6. Animal preparation for SDT
<ol>
	<li>Anesthetize the mouse using an isoflurane-O2&#160;gas mixture in an induction chamber. Set the gas flow rate to 1.0 mL/min and the vaporizer to 2.0% for anesthesia induction, typically requiring 3-5 min in the chamber (see&#160;Table of Materials).</li>
	<li>Assess the animal for adequate sedation by pinching the toe. Apply an ophthalmic ointment to the eyes to avoid dryness of the corneas. 
	NOTE: Monitor the anesthesia depth throughout the procedure.</li>
	<li>Inject the mouse with 40 &#181;L of gadolinium contrast agent through the tail vein (see Table of Materials).</li>
	<li>Remove any hair obstructing the scalp above the skull using a hair removal cream and an electronic shaver.</li>
	<li>Secure the animal to the MRI bed using the following steps (see Figure 3).
	<ol>
		<li>Connect the inlet tube of the MRI bed (Figure 3A) to a source of isoflurane for anesthesia, and set the gas flow rate to 1.0 mL/min and the vaporizer to 2.0% for anesthesia induction. Connect the outlet tube to a charcoal filter canister for anesthesia absorption.</li>
		<li>Place the nose cone piece into its slot, as shown in Figure 3B. Slide the bite bar through the bite bar holes in both the nose cone and at the end of the bed, as shown, with the bite guard end hovering over the open well in the MRI bed.</li>
		<li>Place the animal on the MRI bed with its ears lined up with the stereotactic ear bar holes and latch its teeth through the bite guard to keep it in place. Slide the nose cone forward so that it is over top of the animal&#39;s snout to provide a steady stream of anesthesia.</li>
		<li>Slide the ear bars through the holes on both sides of the MRI bed and raise the animal&#39;s head until both ear bars can fit into the mouse&#39;s ear canals. 
		NOTE: Do not push too far, as this can damage the animal&#39;s ear drums.</li>
		<li>Ensure that the animal is in a comfortable position. Then, using a flathead screwdriver, screw in the MRI-compatible screws to lock in both ear bars, the nose cone, and the bite bar. This will lock the animal into place and prevent any head movement until the animal is removed from the bed. 
		NOTE: Make sure that there is no disturbance to the animal&#39;s positioning on the MRI bed following this step. If the animal moves, then the entire setup (step 6.5) will need to be repeated, regardless of how far along the protocol is.</li>
		<li>While the animal is waiting, place the MRI bed on a warm heat pad to maintain the body temperature. 
		NOTE: Monitor and maintain&#160;the body temperature throughout the procedure.</li>
	</ol>
	</li>
	<li>Isoflurane is continuously supplied to the animal via tubes attached to the nose cones throughout the treatment when the animal bed is fixed on the FUS system, using the fixtures provided on the platform.</li>
</ol>
7. MRI procedures
<ol>
	<li>Take the MRI bed with the animal stereotactically fixed and place it in the MRI cradle previously connected to the MRI scanner (see Table of Materials), by slotting the MRI bed hole at its end to the peg on the MRI cradle, as shown in Figure 3. Attach the inlet and outlet anesthesia tubes to the corresponding tubes in the MRI machine.</li>
	<li>Slide the MRI cradle containing the animal into the MRI bore, making sure to keep the same positioning as where the phantom was placed.</li>
	<li>Perform a localizer to see the location of the animal brain and then a post-contrast T1-weighted MRI scan covering the entire brain using the MRI settings from Table 2.</li>
	<li>Export the MRI scan as a set of uncompressed DICOM files, one for each slice.</li>
</ol>
8. Focused ultrasound treatment (Figure 4)
<ol>
	<li>Remove the animal from the MRI cradle after completion of the scan and place it on the platform by slotting the front of the bed into its corresponding peg at the back of the platform, and slotting the back of the bed into its corresponding peg.</li>
	<li>Connect the inlet tube of the MRI bed to a source of isoflurane for anesthesia and set the gas flow rate to 1.0 mL/min and the vaporizer to 2.0% for maintenance of anesthesia. Connect the outlet tube to a charcoal filter canister for anesthesia absorption.</li>
	<li>Transfer the set of DICOM files to the computer and place them in a folder in the main working directory of the current study. 
	NOTE: Refer to step 5.9 for file requirements.</li>
	<li>In the software, go to the main initialization page and turn on Jog Mode by clicking on Guided Focus Finding and then click on Jog Mode On.</li>
	<li>Place a dab of centrifuged ultrasound gel on the animal&#39;s head, with enough gel to cover the entire scalp above the skull.</li>
	<li>Pour DI and degassed water into the water bath up to 80%, and then slot the water bath onto its corresponding columns on the platform.</li>
	<li>Lower the water bath filled with DI water until the bottom membrane touches the ultrasound gel on the animal&#39;s head, forming a coupling surface between the water and the gel. Make sure that the ultrasound gel covers the entire head of the animal to the water bath and that there are no air bubbles in the ultrasound gel between the water bath and the mouse&#39;s scalp.</li>
	<li>Submerge the ultrasound transducer into the water bath, ensuring no air bubbles are forming on the transducer surface.</li>
	<li>Lower the transducer arm using Jog Mode Down toward the partly submerged transducer, and couple it to the transducer by aligning the magnetic slots with each other while the transducer surface remains submerged.</li>
	<li>Turn off Jog Mode, and then Exit Focused Finding, as done during the initialization step. Make sure to confirm the home position as done before.</li>
	<li>Go to the treatment tab, and then upload the post-contrast T1-weighted MRI files by clicking the folder icon in the top middle of the screen. Select the folder containing the DICOM files that were uploaded to the computer previously and open them. This step should automatically load all DICOM files in the top right panel.</li>
	<li>Next, in the Sonication Mode tab, select the proper treatment mode by choosing either Burst or Continuous Wave. If on burst mode, in the Sonication Settings tab, enter the burst length, burst period, and number of periods. These parameters will correspond to each sonication location. If in continuous wave mode, only enter the sonication duration. 
	NOTE: In this study, continuous wave mode was used for a 120 s duration.</li>
	<li>Click on the target icon in the top middle of the page and select the proper locations on the correct MRI slice where the FUS focal region is to be aimed. A light red square with a red circle will be present where clicked. More than one selection can be chosen.</li>
	<li>Left of the MRI image is a table, which will populate with the 3D coordinates of the focal regions selected. In the last column of the table, enter the power level to be sonicated at each focal spot, which can be calculated separately for each transducer used to obtain the desired pressure or intensity levels. Click on each focal spot in the table to confirm, which will result in the coordinates being highlighted, and the corresponding focal region on the image will turn blue. 
	NOTE: Refer to the transducer datasheet to determine how to translate the power level entered into the table into pressure or intensity, as these values are transducer specific.</li>
	<li>Once satisfied with all focal regions, select Motion Test. This will prompt the software to move the transducer to all of the focal spots to ensure the movements are possible. After confirmation, the software will indicate Motion Test Complete. If an error occurs, adjust the focal regions so that they can fit within the 3D axes of the transducer motors.</li>
	<li>Once ready, select Begin Sonication to begin the sonication protocol, moving the transducer and applying the correct FUS parameters at each focal region selected.</li>
	<li>Once the sonication protocol is finished, uncouple the transducer from the transducer arm, remove the water bath from the platform, and wipe off the ultrasound gel from the animal&#39;s scalp.</li>
	<li>Unscrew the bite and ear bars, remove the animal from the MRI bed, and then place the animal on a warm pad until the animal wakes up from anesthesia. At this point, return the animal to its cage.</li>
	<li>For subsequent animals, repeat the same protocol, starting from section 6.</li>
	<li>Once the desired animals are treated, clean all surfaces touched by animals with 70% ethanol (see Table of Materials).</li>
	<li>Quit out of the software, and then shut down and turn off each piece of equipment.</li>
</ol>
9. Post-treatment steps
<ol>
	<li>Repeat the in vivo imaging system (IVIS) protocol from section 1 for bioluminescence measuring 24 h post-treatment. To analyze treatment effectiveness, compare the bioluminescence recordings before and after treatment to determine the growth rate for both treated and untreated groups.</li>
	<li>Perform follow-up MRI scans using the same settings from step 6.4. Using contrast-enhanced scans, compare the mean greyscale intensity of the tumor region or calculate the total volume that is covered by the contrast agent to determine differences in tumor volume pre- and post-treatment.</li>
</ol>

<ol>
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M., 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=Sonodynamic+therapy+in+combination+with+photodynamic+therapy+shows+enhanced+long-term+cure+of+brain+tumor.">Sonodynamic therapy in combination with photodynamic therapy shows enhanced long-term cure of brain tumor.</a> Scientific Reports. 10 (1), 21791 (2020).</li><li>Wu, S. -. K., Santos, M. A., Marcus, S. L., Hynynen, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MR-guided+focused+ultrasound+facilitates+sonodynamic+therapy+with+5-Aminolevulinic+acid+in+a+rat+glioma+model.">MR-guided focused ultrasound facilitates sonodynamic therapy with 5-Aminolevulinic acid in a rat glioma model.</a> Scientific Reports. 9 (1), 10465 (2019).</li><li>Zhang, Q., 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=Sonodynamic+therapy-assisted+immunotherapy:+A+novel+modality+for+cancer+treatment.">Sonodynamic therapy-assisted immunotherapy: A novel modality for cancer treatment.</a> Cancer Science. 109 (5), 1330-1345 (2018).</li><li>Nonaka, M., 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=Sonodynamic+therapy+consisting+of+focused+ultrasound+and+a+photosensitizer+causes+a+selective+antitumor+effect+in+a+rat+intracranial+glioma+model.">Sonodynamic therapy consisting of focused ultrasound and a photosensitizer causes a selective antitumor effect in a rat intracranial glioma model.</a> Anticancer Research. 29 (3), 943-950 (2009).</li><li>Pi, Z., 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=Sonodynamic+therapy+on+intracranial+glioblastoma+xenografts+using+sinoporphyrin+sodium+delivered+by+ultrasound+with+microbubbles.">Sonodynamic therapy on intracranial glioblastoma xenografts using sinoporphyrin sodium delivered by ultrasound with microbubbles.</a> Annals of Biomedical Engineering. 47 (2), 549-562 (2019).</li><li>Rodgers, G., 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=Virtual+histology+of+an+entire+mouse+brain+from+formalin+fixation+to+paraffin+embedding.+Part+1:+Data+acquisition,+anatomical+feature+segmentation,+tracking+global+volume+and+density+changes.">Virtual histology of an entire mouse brain from formalin fixation to paraffin embedding. 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The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. Amir Manbachi teaches and consults for BK Medical (GE Healthcare), Neurosonics Medical, and is an inventor on a number of patent-pending FUS technologies. Betty Tyler has research funding from NIH and is a co-owner of Accelerating Combination Therapies*. Ashvattha Therapeutics Inc. has also licensed one of her patents, and she is a stockholder for Peabody Pharmaceuticals (*includes equity or options).

New therapeutic and efficacious treatment options are necessary for patients with GBM. This protocol has outlined a preclinical FUS-mediated treatment for GBM that is currently undergoing extensive investigation for clinical translation. Although SDT has exciting potential, there is still much to understand and optimize in the preclinical setting.
One of the most important components of this protocol is utilizing MR-guided FUS to target the tumor for maximal efficacy. Using a phantom, a 3D coordinate space can be created, where each pixel of axial MRI slices can be assigned a coordinate. Then, a simple procedure of selecting the sonication location on the MR image informs the transducer where to aim. The preclinical FUS system used is highly versatile and applicable when needing to target locations of specific pathology such as a tumor, including deeper seated tumors which would be hard to target without imaging confirmation. Using gadolinium as a contrast agent, there is clear visualization of the tumor, allowing the user to make informed decisions when choosing targets. The advantage that SDT has over many other treatments is that it is a tumor-specific therapy. Low-intensity FUS should only target the tumor tissue, while leaving the healthy brain parenchyma relatively untouched3,8.
The results of this experiment highlight how the advantages of this protocol can lead to therapeutic results that are similar to other findings in the literature for SDT. Figure 5 shows that within as little as 24 h following the day of treatment, there was a slowdown of tumor growth in the treated cohort. Although insignificant using this small sample size, significance might result with a larger number of animals. This delay in tumor growth is similar to what was shown in the pioneering paper on this subject by Wu et al. (2019), which exhibited slowed tumor growth over time in treated animals, as well as increased survival times9.
Considerations that were made when designing this protocol included animal strain, tumor type, and sonosensitizing agent selection. Athymic nude mice were chosen for this protocol for multiple reasons. First, the nude mouse is easier to sonicate as the lack of hair prevents any attenuation. Also, the lack of an immune system allows for the implantation of patient-derived xenografts (PDXs) so that the tumor model more closely resembles the clinical situation. The downside of using an athymic model is that the immune system cannot be characterized, so any SDT-generated immune response will not be measured in these studies10. The tumor line chosen is an aggressive and fast-growing PDX line. The time of treatment is very important because the establishment of the tumor must be verified, but the tumor burden should not fill the cranial hemisphere. Different cell lines require different incubation times to achieve an optimally sized tumor for preclinical experimentation. In this protocol, 5-ALA was used as the sonosensitizer because of its preferential uptake in GBM tumors, which has been confirmed in vitro for this cell line in previous experiments (unpublished data). Other sonosensitizers can be substituted and tested to determine the compound most suitable for efficacy and safety. Finally, treatment was commenced 3 h post 5-ALA injection, as previous literature has shown that this is the optimal time with that injection dosage5.
The FUS parameters chosen in this protocol (10 W/cm2 for 2 min at 515 kHz at each target location) were decided based on a review of previous literature and initial experiments4,9. A grid of sonication points covering the entire tumor was chosen in order to generate the ROS effect throughout the entire tumor. The intensity used here is higher than other publications, but at a short time span, this is not expected to lead to any adverse temperature-related effects, as intensities up to 25 W/cm2 have been successfully used in a mouse model without significant side effects11. Importantly, no standardized or optimized set of FUS parameters has been published in the literature. Therefore, the specific values that are reported here can be adjusted to determine the optimal set of parameters, leading to the maximal reduction of tumor tissue while maintaining safety. Additionally, as different cell lines have varying levels of vascularization and hypoxia, this treatment may need to be adjusted. We have shown overall decreased tumor growth (Figure 5) within 24 h of SDT treatment, although the parameters need to be optimized and more animals need to be tested to determine the maximal effect of this treatment. Post-treatment MRI scans show no appearance of lesions created by FUS treatment in healthy tissue, with the effect localized to tumor tissue (Figure 6). There is also the opportunity to combine SDT with other FUS techniques, such as transiently permeabilizing the blood-brain barrier, to maximize 5-ALA uptake in the tumor12. This protocol can be further supplemented by performing various histology techniques to check for safety and efficacy at the structural level. A hematoxylin and eosin (H&#38;E) stain can be performed to check for structural or tumor damage13, while a terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) stain can be performed to check for cellular apoptosis14. Regardless, this protocol presents a safe and tumor-specific treatment where changes are noticeable even 24 h post-treatment, which is apparent by comparing the growth rate of tumors treated with SDT and untreated tumors, as well as comparing tumor slices before and after sonication.
With any protocol, there are always disadvantages or limitations that need to be weighed. The main limitation of the current protocol is time and expense. Meanwhile, one of the advantages of this protocol is its automated focused aim. To accomplish this focused procedure, MRI scans need to be taken for each individual animal to ensure that the targeting of the tumor is correct, a process which can be both time-consuming and expensive. Additionally, depending on the number of focal spots desired, the amount of time to perform this protocol could be hours for even just a few animals, resulting in low experimental animal numbers. Despite these drawbacks, this targeted noninvasive protocol remains a feasible preference when compared to open surgery options.
In conclusion, this protocol showed the ability of SDT treatment to decrease tumor growth in the brain within 24 h of treatment while maintaining healthy neural tissue in a preclinical mouse model. Studies of the effectiveness of SDT and optimizing the various parameters to increase ROS production are necessary to make this treatment clinically suitable. New avenues should be explored for the use of SDT as a noninvasive therapeutic model.

Glioblastoma (GBM) is a form of highly aggressive brain cancer that has an incidence of 3.21 per 100,000 people and is the most common malignant brain tumor1. The current standard of care includes surgical resection, radiation, and chemotherapy2. Due to the tumor&#39;s invasive and infiltrative nature, complete tumor resection is rare. Residual tissue at the tumor margins results in a high rate of tumor recurrence and a low survival rate of less than 6% after 5 years1.
Due to this prognosis, researchers are exploring new therapeutic options to combat this deadly disease. Sonodynamic therapy (SDT) is a noninvasive treatment that combines low-intensity focused ultrasound (FUS) and sonosensitizing agents to produce a cytotoxic effect in targeted cells3. As an example, porphyrin-based sonosensitizers such as 5-aminolevulinic acid (5-ALA) are preferentially taken up by tumor cells and increase reactive oxidative species (ROS) production to damaging levels when exposed to focused ultrasound. Overexpressed levels of ROS&#160;in cells can damage cellular structures and trigger apoptosis. Since 5-ALA is preferentially taken up by tumor cells, healthy tissue within the treatment region is unharmed3,4. Preliminary in vitro studies have revealed that many cancer cells are lysed by SDT treatment, though the rate of cell death is dependent on the cell line. Preliminary in vivo studies yield similar results, confirming that SDT can trigger apoptosis5.
This protocol aims to describe effective techniques and parameters for the SDT treatment of rodent models with intracranially implanted GBM cells using a benchtop FUS research platform. Researchers can use this protocol to perform and optimize SDT for translational FUS research.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5% Trypsin-EDTA</td>
			<td>Thermo Fisher Scientific</td>
			<td>15400054</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL Syringes</td>
			<td>BD</td>
			<td>309597</td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#181;L Hamilton syringe</td>
			<td>Hamilton Company</td>
			<td>49AL65</td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#181;L Pipette tips</td>
			<td>USAScientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1000 mL Flask</td>
			<td>Corning</td>
			<td>MP-34514-25</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL conical tubes</td>
			<td>Corning</td>
			<td>CLS430791</td>
			<td></td>
		</tr>
		<tr>
			<td>200 Proof ethanol</td>
			<td>PharmCo</td>
			<td>111000200</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL pipettes</td>
			<td>Falcon</td>
			<td>357543</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL Conical tubes</td>
			<td>Corning</td>
			<td>430290</td>
			<td></td>
		</tr>
		<tr>
			<td>500 mL filter</td>
			<td>Corning</td>
			<td>431097</td>
			<td></td>
		</tr>
		<tr>
			<td>5-Aminolevulinic acid hydrochloride&#160;</td>
			<td>Research Products International</td>
			<td>A11250</td>
			<td></td>
		</tr>
		<tr>
			<td>7T PET-MR system</td>
			<td>Bruker</td>
			<td>Biospec 70/30</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum foil</td>
			<td>Reynolds Brand</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Amplifier</td>
			<td>FUS Instruments</td>
			<td>2175</td>
			<td></td>
		</tr>
		<tr>
			<td>Athymic nude mice</td>
			<td>Charles River Laboratories</td>
			<td>Strain Code 490</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone drill</td>
			<td>Foredom</td>
			<td>HP4-917</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Thermo Fisher Scientific</td>
			<td>75004261</td>
			<td></td>
		</tr>
		<tr>
			<td>Charcoal isoflourane waste container</td>
			<td>Patterson scientific</td>
			<td>78909457</td>
			<td></td>
		</tr>
		<tr>
			<td>Computer</td>
			<td>FUS Instruments</td>
			<td>2269</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover glass</td>
			<td>Fisherbrand</td>
			<td>12-545J</td>
			<td></td>
		</tr>
		<tr>
			<td>Desktop monitor</td>
			<td>ASUS</td>
			<td>VZ239H</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Luciferin</td>
			<td>Gold Biotechnology</td>
			<td>LUCK-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Thermo Fisher Scientific</td>
			<td>11965092</td>
			<td></td>
		</tr>
		<tr>
			<td>Electronic shaver</td>
			<td>Wahl&#160;</td>
			<td>93235-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf tubes</td>
			<td>Posi-Click</td>
			<td>1149K01</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum</td>
			<td>Thermo Fisher Scientific</td>
			<td>16000044</td>
			<td></td>
		</tr>
		<tr>
			<td>Formalin</td>
			<td>Thermo Fisher Scientific</td>
			<td>SF100-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Function generator</td>
			<td>Siglent</td>
			<td>QS0201X-E01B</td>
			<td></td>
		</tr>
		<tr>
			<td>Gadolinium contrast agent (Gadavist)</td>
			<td>McKesson Corporation</td>
			<td>2068062</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze</td>
			<td>Henry Schein</td>
			<td>101-4336</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat lamp</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Heat pad</td>
			<td>Kent Scientific</td>
			<td>RT-0501</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemocytometer</td>
			<td>Electron Microscopy Sciences</td>
			<td>63514-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Induction chamber</td>
			<td>Patterson scientific</td>
			<td>78933388</td>
			<td></td>
		</tr>
		<tr>
			<td>Isofluorane vaporizer</td>
			<td>Patterson scientific</td>
			<td>78916954</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane&#160;</td>
			<td>Covetrus</td>
			<td>29405</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane system</td>
			<td>Patterson Scientific</td>
			<td>78935903</td>
			<td></td>
		</tr>
		<tr>
			<td>IVIS spectrum</td>
			<td>Perkin Elmer</td>
			<td>124262</td>
			<td></td>
		</tr>
		<tr>
			<td>Lightfield microscope</td>
			<td>BioTek</td>
			<td>Cytation 5</td>
			<td></td>
		</tr>
		<tr>
			<td>Nair</td>
			<td>Church and Dwight Co.</td>
			<td>42010553</td>
			<td></td>
		</tr>
		<tr>
			<td>Ophthalmic ointment&#160;</td>
			<td>Puralube vet ointment</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>P-20 pippette</td>
			<td>Rainin</td>
			<td>17008650</td>
			<td></td>
		</tr>
		<tr>
			<td>Patient derived xenographs</td>
			<td>Mayo Clinic</td>
			<td>M59</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomyosin</td>
			<td>Thermo Fisher Scientific</td>
			<td>10378016</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline</td>
			<td>Thermo Fisher Scientific</td>
			<td>70-011-069</td>
			<td></td>
		</tr>
		<tr>
			<td>Pippetter</td>
			<td>Drummond</td>
			<td>4-000-101</td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone-iodine</td>
			<td>Covetrus</td>
			<td>PI050CV</td>
			<td></td>
		</tr>
		<tr>
			<td>RK-50 MRgFUS system&#160;</td>
			<td>FUS Instruments</td>
			<td>2182</td>
			<td></td>
		</tr>
		<tr>
			<td>Scale</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td>Covetrus</td>
			<td>7319</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel handle</td>
			<td>Fine Science Tools</td>
			<td>91003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Screwdriver set</td>
			<td>Jakemy</td>
			<td>JM-8160</td>
			<td></td>
		</tr>
		<tr>
			<td>Skin marker</td>
			<td>Time Out</td>
			<td>D538,851</td>
			<td></td>
		</tr>
		<tr>
			<td>Staple remover</td>
			<td>MikRon</td>
			<td>ACR9MM</td>
			<td></td>
		</tr>
		<tr>
			<td>Stapler</td>
			<td>MikRon</td>
			<td>ACA9MM</td>
			<td></td>
		</tr>
		<tr>
			<td>Staples</td>
			<td>Clay Adams</td>
			<td>427631</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotactic frame</td>
			<td>Kopf Instruments</td>
			<td>5000</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotactic MRI prototype plastic imaging fixture</td>
			<td>FUS Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>T-25 culture flask</td>
			<td>Corning</td>
			<td>430641U</td>
			<td></td>
		</tr>
		<tr>
			<td>Transducer and matching box</td>
			<td>FUS Instruments</td>
			<td>T515H750-118</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic degasser</td>
			<td>FUS Instruments</td>
			<td>2259</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasound gel</td>
			<td>ParkerLabs</td>
			<td>&#160;01-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>FUS Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Covetrus</td>
			<td>1XYL006</td>
			<td></td>
		</tr>
	</tbody>
</table>

sonodynamic therapy for the treatment of glioblastoma multiforme in a mouse model using a portable benchtop focused ultrasound system

Sonodynamic therapy (SDT) is an application of focused ultrasound (FUS) that enables a sonosensitizing agent to prime tumors for increased sensitivity during sonication. Unfortunately, current clinical treatments for glioblastoma (GBM) are lacking, leading to low long-term survival rates among patients. SDT is a promising method for treating GBM in an effective, noninvasive, and tumor-specific manner. Sonosensitizers preferentially enter tumor cells compared to the surrounding brain parenchyma. The application of FUS in the presence of a sonosensitizing agent generates reactive oxidative species resulting in apoptosis. Although this therapy has been shown previously to be effective in preclinical studies, there is a lack of established standardized parameters. Standardized methods are necessary to optimize this therapeutic strategy for preclinical and clinical use. In this paper, we detail the protocol to perform SDT in a preclinical GBM rodent model using magnetic resonance-guided FUS (MRgFUS). MRgFUS is an important feature of this protocol, as it allows for specific targeting of a brain tumor without the need for invasive surgeries (e.g., craniotomy). The benchtop device used here can focus on a specific location in three dimensions by clicking on a target on an MRI image, making target selection a straightforward process. This protocol will provide researchers with a standardized preclinical method for MRgFUS SDT, with the added flexibility to change and optimize parameters for translational research.

Tumor size declines in animals treated with SDT 24 h post-treatment. 
On the day of SDT treatment, the original average bioluminescence signal for the control and treatment groups (N = 4 each) was 2.0 x 106 &#177; 3.1 x 106 and 2.3 x 106 &#177; 1.3 x 106 p/s/cm2/sr, respectively. The average bioluminescence values corresponding to tumor size before treatment between the two groups were not statistically significant (p = 0.89). The average bioluminescence signal of the treatment group was 3.57 x 106 &#177; 2.3 x 106 24 h following SDT, while the bioluminescent signal of the control group was increased to 5.5 x 106 &#177; 8.2 x 106 p/s/cm2/sr. As shown in Figure 5, this corresponds to a growth rate of 83.4% &#177; 78% and 172% &#177; 34%, respectively, assuming exponential growth (p = 0.08). Of the four treated animals, three had lower growth rates post-treatment compared to controls. There was one outlier in the treatment group that showed comparable growth to controls, skewing the deviations.
Additionally, the animals underwent subsequent contrast-enhanced MR imaging the day following treatment for pre- and post-treatment comparison of the tumor. The average grayscale intensity of contrast agent in tumors was conducted across each MRI slice for each animal to measure how much contrast agent was entering tumors following treatment, as an estimate of tumor size. Pre-treatment, the average tumor grayscale intensity between control and treated groups was similar. On average, this greyscale intensity increased in the control group to a larger magnitude than in treated groups, although this was not significant (p = 0.47). This data can be seen in Table 2. The high variability of these results is potentially due to the fact that MRIs were taken only 24 h post-treatment, at which time the therapeutic potential of SDT is only beginning to occur. Even so, Figure 6 shows an example of the lesions created by SDT.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65114/65114fig01.jpg" /> 
Figure 1: FUS system setup. (Top) MRgFUS System with labeled components. (Front) 1. Platform. 2. Axis motorized transducer arm. 3. FUS transducer. 4. Water bath. 5. MRI bed. 6. Monitor. 7. Transducer impedance matching box. 8. Power amplifier box. 9. Function generator. 10. Function generator channel 1 BNC port. 11. Desktop computer. (Bottom) MRgFUS system with color-coded wiring schematic with the following connections. (Back) 1. Power cords. 2. Monitor HDMI to desktop HDMI. 3. Port B USB B to desktop USB A. 4. Oscilloscope LAN ethernet to desktop ethernet. 5. Motion interface ethernet to desktop ethernet. 6. Oscilloscope aux in/out BNC to AWG input BNC. 7. Oscilloscope channel 1 (Front) BNC to SYNC BNC. 8. Matching box output BNC to RF input coaxial. 9. RF output coaxial to matching box coaxial. <a href="https://www.jove.com/files/ftp_upload/65114/65114fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65114/65114fig02.jpg" /> 
Figure 2: Phantom registration. (A)&#160;System setup and software during phantom registration. (B) Screenshot of the phantom registration screen, where the red circle is the selected circumference of the axial cross section. (C)&#160;Phantom placed on the MRI bed, top view. (D)&#160;Side view of the phantom, where the red line is in the axial slice corresponding to the circle in C. <a href="https://www.jove.com/files/ftp_upload/65114/65114fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65114/65114fig03.jpg" /> 
Figure 3: Animal placement. (A) MRI bed and cradle, with various parts labeled: 1. MRI cradle. 2. Stereotactic MRI bed. 3. Ear bars. 4. Bite bar. 5. Nose cone. 6. MRI bed peg hole. 7. Isoflurane anesthesia tubes. (B) Illustration representing the placement of the mouse on the MRI bed and placement on the cradle, with the RF coil (Orange) (illustration modified using Biorender 2022 template). (C) Illustration representing the placement of the mouse on the MRI bed during a FUS treatment (illustration modified using Biorender 2022 template). <a href="https://www.jove.com/files/ftp_upload/65114/65114fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65114/65114fig04.jpg" /> 
Figure 4: Focal point selection. Example of a sonication point selection in a single animal. Each column represents a T1-weighted post-contrast MRI slice where each slice is 0.5 mm in the proximal (slice 1) to distal (slice 5) direction. The tumor boundary was manually segmented and is outlined in red (row 1), and the corresponding sonication locations (row 2) are represented by a light green square (focal max center) and blue circle (half max focal circumference). Each location was sonicated for 2 min. <a href="https://www.jove.com/files/ftp_upload/65114/65114fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65114/65114fig05.jpg" /> 
Figure 5: Growth rate following SDT. The growth rate of GBM tumors from pre- to 24 h post-SDT treatment in treated and untreated (control) animals with intracranial M59 tumors based on the measured luminescence. Error bars indicate standard deviation. A two-sample student&#39;s T-test was performed to determine significance. <a href="https://www.jove.com/files/ftp_upload/65114/65114fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65114/65114fig06.jpg" /> 
Figure 6: SDT generated lesion. Pre- and post-contrast enhance T1-weighted MRI scans from an animal model featuring a representative axial slice showing a lesion in the tumor created by SDT. (Left) MRI scan taken prior to SDT treatment, with the tumor outlined in red. (Middle) FUS focal point selection where the maximum pressure is represented by light blue circles and the half-maximum pressure regions are represented by blue circles. (Right) Post-SDT MRI scans, where the tumor is outlined in red. SDT-created lesions and the syringe hole for implantation are shown. <a href="https://www.jove.com/files/ftp_upload/65114/65114fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sequence</td>
			<td>T1</td>
		</tr>
		<tr>
			<td>Repetition time</td>
			<td>3000 ms</td>
		</tr>
		<tr>
			<td>Echo time</td>
			<td>30 ms</td>
		</tr>
		<tr>
			<td>Slice thickness</td>
			<td>0.5 mm</td>
		</tr>
		<tr>
			<td>Number of slices</td>
			<td>25</td>
		</tr>
		<tr>
			<td>Pixel spacing</td>
			<td>0.187 mm x 0.187 mm</td>
		</tr>
		<tr>
			<td>Acquisition matrix</td>
			<td>133 x 133</td>
		</tr>
		<tr>
			<td>Averages</td>
			<td>4</td>
		</tr>
	</tbody>
</table>
Table 1: MRI settings.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Control</td>
			<td>SDT Group</td>
			<td>P-Value</td>
		</tr>
		<tr>
			<td>Pre-Treatment</td>
			<td>7.49 x 103 &#177; 2.2 x 103</td>
			<td>7.48 x 103 &#177; 1 x 103</td>
			<td>0.99</td>
		</tr>
		<tr>
			<td>Post-Treatment</td>
			<td>8.79 x 103 &#177; 7.7 x 102&#160;</td>
			<td>7.95 x 103 &#177; 1.1 x 103&#160;</td>
			<td>0.33</td>
		</tr>
		<tr>
			<td>Percent Difference</td>
			<td>16% &#177; 16%</td>
			<td>7% &#177; 12%&#160;</td>
			<td>0.47</td>
		</tr>
	</tbody>
</table>
Table 2: Post-contrast enhanced T1-weighted MRI greyscale.

Watch this Scientific Journal Video about Sonodynamic Therapy for the Treatment of Glioblastoma Multiforme in a Mouse Model Using a Portable Benchtop Focused Ultrasound System at JoVE.com

Sonodynamic Therapy for the Treatment of Glioblastoma Multiforme in a Mouse Model Using a Portable Benchtop Focused Ultrasound System

Here, we describe a protocol that details how to perform sonodynamic therapy in an in vivo mouse glioblastoma model using magnetic resonance-guided focused ultrasound.

Sonodynamic therapy (SDT) is an application of focused ultrasound (FUS) that enables a sonosensitizing agent to prime tumors for increased sensitivity ...

sonodynamic-therapy-for-treatment-glioblastoma-multiforme-mouse-model

Research

JoVE Journal

Bioengineering

2.3K Views.  Johns Hopkins University School of Medicine. Here, we describe a protocol that details how to perform sonodynamic therapy in an in vivo mouse glioblastoma model using magnetic resonance-guided focused ultrasound. 

Video: Sonodynamic Therapy for the Treatment of Glioblastoma Multiforme in a Mouse Model Using a Portable Benchtop Focused Ultrasound System

A Method for Ovarian Follicle Encapsulation and Culture in a Proteolytically Degradable 3 Dimensional System

The ovarian follicle is the functional unit of the ovary that secretes sex hormones and supports oocyte maturation. In vitro follicle techniques provide a tool to model follicle development in order to investigate basic biology, and are further being developed as a technique to preserve fertility in the clinic1-4. Our in vitro culture system employs hydrogels in order to mimic the native ovarian environment by maintaining the 3D follicular architecture, cell-cell interactions and paracrine signaling that direct follicle development 5. Previously, follicles were successfully cultured in alginate, an inert algae-derived polysaccharide that undergoes gelation with calcium ions6-8. Alginate hydrogels formed at a concentration of 0.25% w/v were the most permissive for follicle culture, and retained the highest developmental competence 9. Alginate hydrogels are not degradable, thus an increase in the follicle diameter results in a compressive force on the follicle that can impact follicle growth10. We subsequently developed a culture system based on a fibrin-alginate interpenetrating network (FA-IPN), in which a mixture of fibrin and alginate are gelled simultaneously. This combination provides a dynamic mechanical environment because both components contribute to matrix rigidity initially; however, proteases secreted by the growing follicle degrade fibrin in the matrix leaving only alginate to provide support. With the IPN, the alginate content can be reduced below 0.25%, which is not possible with alginate alone 5. Thus, as the follicle expands, it will experience a reduced compressive force due to the reduced solids content. Herein, we describe an encapsulation method and an in vitro culture system for ovarian follicles within a FA-IPN. The dynamic mechanical environment mimics the natural ovarian environment in which small follicles reside in a rigid cortex and move to a more permissive medulla as they increase in size11. The degradable component may be particularly critical for clinical translation in order to support the greater than 106-fold increase in volume that human follicles normally undergo in vivo .

A new method for ovarian follicle encapsulation in a 3D fibrin-alginate interpenetrating network is described. This system combines structural support with proteolytic degradation to support the development of immature follicles to produce mature oocytes. This method may be applied to culture cell aggregates to maintain cell-cell contacts without limiting expansion.

The ovarian follicle is the functional unit of the ovary that secretes sex hormones and supports oocyte maturation. In vitro  follicle techniques provide ...

Anatomical Reconstructions of the Human Cardiac Venous System using Contrast-computed Tomography of Perfusion-fixed Specimens

A detailed understanding of the complexity and relative variability within the human cardiac venous system is crucial for the development of cardiac devices that require access to these vessels. For example, cardiac venous anatomy is known to be one of the key limitations for the proper delivery of cardiac resynchronization therapy (CRT)1 Therefore, the development of a database of anatomical parameters for human cardiac venous systems can aid in the design of CRT delivery devices to overcome such a limitation. In this research project, the anatomical parameters were obtained from 3D reconstructions of the venous system using contrast-computed tomography (CT) imaging and modeling software (Materialise, Leuven, Belgium). The following parameters were assessed for each vein: arc length, tortuousity, branching angle, distance to the coronary sinus ostium, and vessel diameter.
	CRT is a potential treatment for patients with electromechanical dyssynchrony. Approximately 10-20% of heart failure patients may benefit from CRT2. Electromechanical dyssynchrony implies that parts of the myocardium activate and contract earlier or later than the normal conduction pathway of the heart. In CRT, dyssynchronous areas of the myocardium are treated with electrical stimulation. CRT pacing typically involves pacing leads that stimulate the right atrium (RA), right ventricle (RV), and left ventricle (LV) to produce more resynchronized rhythms. The LV lead is typically implanted within a cardiac vein, with the aim to overlay it within the site of latest myocardial activation. 
	We believe that the models obtained and the analyses thereof will promote the anatomical education for patients, students, clinicians, and medical device designers. The methodologies employed here can also be utilized to study other anatomical features of our human heart specimens, such as the coronary arteries. To further encourage the educational value of this research, we have shared the venous models on our free access website: <a href="http://www.vhlab.umn.edu/atlas" target="_blank">www.vhlab.umn.edu/atlas</a>.

The objective of this research is to recreate and then access the anatomy of the human cardiac venous system using 3D reconstructions generated from contrast-computed tomography scans.

A detailed  understanding of the complexity and relative variability within the human  cardiac venous system is crucial for the development of cardiac ...

Operation of a Benchtop Bioreactor

Fermentation systems are used to provide an optimal growth environment for many different types of cell cultures. The ability afforded by fermentors to carefully control temperature, pH, and dissolved oxygen concentrations in particular makes them essential to efficient large scale growth and expression of fermentation products. This video will briefly describe the advantages of the fermentor over the shake flask. It will also identify key components of a typical benchtop fermentation system and give basic instruction on setup of the vessel and calibration of its probes. The viewer will be familiarized with the sterilization process and shown how to inoculate the growth medium in the vessel with culture. Basic concepts of operation, sampling, and harvesting will also be demonstrated. Simple data analysis and system cleanup will also be discussed.

Fermentors are used to increase culture yield and productivity of bioengineered cells. After screening multiple microbial or animal cell culture candidates in shake flasks, the next logical step is to increase the selected culture&#x2019;s biomass with the fermentor. This video demonstrates the setup and operation of a typical benchtop bioreactor system.

Fermentation systems are used to provide an optimal growth environment for many different types of cell cultures. The ability afforded by fermentors to ...

Real-time Monitoring of High Intensity Focused Ultrasound (HIFU) Ablation of In Vitro Canine Livers Using Harmonic Motion Imaging for Focused Ultrasound (HMIFU)

Harmonic Motion Imaging for Focused Ultrasound (HMIFU) is a technique that can perform and monitor high-intensity focused ultrasound (HIFU) ablation. An oscillatory motion is generated at the focus of a 93-element and 4.5 MHz center frequency HIFU transducer by applying a 25 Hz amplitude-modulated signal using a function generator. A 64-element and 2.5 MHz imaging transducer with 68kPa peak pressure is confocally placed at the center of the HIFU transducer to acquire the radio-frequency (RF) channel data. In this protocol, real-time monitoring of thermal ablation using HIFU with an acoustic power of 7 W on canine livers in vitro is described. HIFU treatment is applied on the tissue during 2 min and the ablated region is imaged in real-time using diverging or plane wave imaging up to 1,000 frames/second. The matrix of RF channel data is multiplied by a sparse matrix for image reconstruction. The reconstructed field of view is of 90&#176; for diverging wave and 20 mm for plane wave imaging and the data are sampled at 80 MHz. The reconstruction is performed on a Graphical Processing Unit (GPU) in order to image in real-time at a 4.5 display frame rate. 1-D normalized cross-correlation of the reconstructed RF data is used to estimate axial displacements in the focal region. The magnitude of the peak-to-peak displacement at the focal depth decreases during the thermal ablation which denotes stiffening of the tissue due to the formation of a lesion. The displacement signal-to-noise ratio (SNRd) at the focal area for plane wave was 1.4 times higher than for diverging wave showing that plane wave imaging appears to produce better displacement maps quality for HMIFU than diverging wave imaging.

Real-time Monitoring of High Intensity Focused Ultrasound HIFU Ablation of In Vitro Canine Livers Using Harmonic Motion Imaging for Focused Ultrasound HMIFU

This article describes real-time monitoring of HIFU ablation in canine liver with high frame rate ultrasound imaging using diverging and plane wave imaging. Harmonic Motion Imaging for Focused Ultrasound is used to image the decrease of acoustic radiation force induced displacement in the ablated region.

Harmonic Motion Imaging for Focused Ultrasound (HMIFU) is a technique that can perform and monitor high-intensity focused ultrasound (HIFU) ablation. An ...

Three-Dimensionally Printed Microfluidic Cross-flow System for Ultrafiltration/Nanofiltration Membrane Performance Testing

Minimization and management of membrane fouling is a formidable challenge in diverse industrial processes and other practices that utilize membrane technology. Understanding the fouling process could lead to optimization and higher efficiency of membrane based filtration. Here we show the design and fabrication of an automated three-dimensionally (3-D) printed microfluidic cross-flow filtration system that can test up to 4 membranes in parallel. The microfluidic cells were printed using multi-material photopolymer 3-D printing technology, which used a transparent hard polymer for the microfluidic cell body and incorporated a thin rubber-like polymer layer, which prevents leakages during operation. The performance of ultrafiltration (UF), and nanofiltration (NF) membranes were tested and membrane fouling could be observed with a model foulant bovine serum albumin (BSA). Feed solutions containing BSA showed flux decline of the membrane. This protocol may be extended to measure fouling or biofouling with many other organic, inorganic or microbial containing solutions. The microfluidic design is especially advantageous for testing materials that are costly or only available in small quantities, for example polysaccharides, proteins, or lipids due to the small surface area of the membrane being tested. This modular system may also be easily expanded for high throughput testing of membranes.&#160;

Design and fabrication of a three-dimensionally (3-D) printed microfluidic cross-flow filtration system is demonstrated. The system is used to test performance and observe fouling of ultrafiltration and nanofiltration (thin film composite) membranes.

Minimization and management of membrane fouling is a formidable challenge in diverse industrial processes and other practices that utilize membrane ...

Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional tissue. Tissue engineering holds great promise as a potential solution with its ability to integrate cells and signaling molecules into a 3-dimensional scaffold. Recent work with tissue engineered tracheal grafts (TETGs) has seen some success but their translation has been limited by graft stenosis, graft collapse, and delayed epithelialization. In order to investigate the mechanisms driving these issues, we have developed a mouse model for tissue engineered tracheal graft implantation. TETGs were constructed using electrospun polymers polyethylene terephthalate (PET) and polyurethane (PU) in a mixture of PET and PU (20:80 percent weight). Scaffolds were then seeded using bone marrow mononuclear cells isolated from 6-8 week-old C57BL/6 mice by gradient centrifugation. Ten million cells per graft were seeded onto the lumen of the scaffold and allowed to incubate overnight before implantation between the third and seventh tracheal rings. These grafts were able to recapitulate the findings of stenosis and delayed epithelialization as demonstrated by histological analysis and lack of Keratin 5 and Keratin 14 basal epithelial cells on immunofluorescence. This model will serve as a tool for investigating cellular and molecular mechanisms involved in host remodeling.

Graft stenosis poses a critical obstacle in tissue engineered airway replacement. To investigate cellular mechanisms underlying stenosis, we utilize a murine model of tissue engineered tracheal replacement with seeded bone marrow mononuclear cells (BM-MNC). Here, we detail our protocol, including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation.

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional ...

Focused Ion Beam Lithography to Etch Nano-architectures into Microelectrodes

With advances in electronics and fabrication technology, intracortical microelectrodes have undergone substantial improvements enabling the production of sophisticated microelectrodes with greater resolution and expanded capabilities. The progress in fabrication technology has supported the development of biomimetic electrodes, which aim to seamlessly integrate into the brain parenchyma, reduce the neuroinflammatory response observed after electrode insertion and improve the quality and longevity of electrophysiological recordings. Here we describe a protocol to employ a biomimetic approach recently classified as nano-architecture. The use of focused ion beam lithography (FIB) was utilized in this protocol to etch specific nano-architecture features into the surface of non-functional and functional single shank intracortical microelectrodes. Etching nano-architectures into the electrode surface indicated possible improvements of biocompatibility and functionality of the implanted device. One of the benefits of using FIB is the ability to etch on manufactured devices, as opposed to during the fabrication of the device, facilitating boundless possibilities to modify numerous medical devices post-manufacturing. The protocol presented herein can be optimized for various material types, nano-architecture features, and types of devices. Augmenting the surface of implanted medical devices can improve the device performance and integration into the tissue.

We have shown that the etching of nano-architecture into intracortical microelectrode devices may reduce the inflammatory response and has the potential to improve electrophysiological recordings. The methods described herein outline an approach to etch nano-architectures into the surface of non-functional and functional single shank silicon intracortical microelectrodes.

With advances in electronics and fabrication technology, intracortical microelectrodes have undergone substantial improvements enabling the production of ...

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of cellular function. The ECM plays a particularly important role in adipose tissue function in obesity, and alterations in adipose tissue ECM deposition and composition are associated with metabolic disease in mice and humans. Tractable in vitro models that permit dissection of the roles of the ECM and cells in contributing to global tissue phenotype are sparse. We describe a novel 3D in vitro model of human ECM-adipocyte culture that permits study of the specific roles of the ECM and adipocytes in regulating adipose tissue metabolic phenotype. Human adipose tissue is decellularized to isolate ECM, which is subsequently repopulated with preadipocytes that are then differentiated within the ECM into mature adipocytes. This method creates ECM-adipocyte constructs that are metabolically active and retain characteristics of the tissues and patients from which they are derived. We have used this system to demonstrate disease-specific ECM-adipocyte crosstalk in human adipose tissue. This culture model provides a tool for dissecting the roles of the ECM and adipocytes in contributing to global adipose tissue metabolic phenotype and permits study of the role of the ECM in regulating adipose tissue homeostasis.

We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of ...

In Vitro Model of Human Cutaneous Hypertrophic Scarring using Macromolecular Crowding

It has been shown that in vivo tissues are highly crowded by proteins, nucleic acids, ribonucleoproteins, polysaccharides, etc. The following protocol applies a macromolecular crowding (MMC) technique to mimic this physiological crowding through the addition of neutral polymers (crowders) to cell cultures in vitro. Previous studies using Ficoll or dextran as crowders demonstrate that the expression of collagen I and fibronectin in WI38 and WS-1 cell lines are significantly enhanced using the MMC technique. However, this technique has not been validated in primary hypertrophic scar-derived human skin fibroblasts (hHSFs). As hypertrophic scarring arises from the excessive deposition of collagen, this protocol aims to construct a collagen-rich in vitro hypertrophic scar model by applying the MMC technique with hHSFs. This optimized MMC model has been shown to possess more similarities with in vivo scar tissue compared to traditional 2-dimensional (2-D) cell culture systems. In addition, it is cost-effective, time-efficient, and ethically desirable compared to animal models. Therefore, the optimized model reported here offers an advanced &#8220;in vivo-like&#8221; model for hypertrophic scar-related studies.

This protocol describes the use of macromolecular crowding to create an in vitro human hypertrophic scar tissue model that resembles in vivo conditions. When cultivated in a crowded macromolecular environment, human skin fibroblasts exhibit phenotypes, biochemistry, physiology, and functional characteristics resembling scar tissue.

It has been shown that in vivo tissues are highly crowded by proteins, nucleic acids, ribonucleoproteins, polysaccharides, etc. The following protocol ...

An In vitro System to Gauge the Thrombolytic Efficacy of Histotripsy and a Lytic Drug

Deep vein thrombosis (DVT) is a global health concern. The primary approach to achieve vessel recanalization for critical obstructions is catheter-directed thrombolytics (CDT). To mitigate caustic side effects and the long treatment time associated with CDT, adjuvant and alternative approaches are under development. One such approach is histotripsy, a focused ultrasound therapy to ablate tissue via bubble cloud nucleation. Pre-clinical studies have demonstrated strong synergy between histotripsy and thrombolytics for clot degradation. This report outlines a benchtop method to assess the efficacy of histotripsy-aided thrombolytic therapy, or lysotripsy.
Clots manufactured from fresh human venous blood were introduced into a flow channel whose dimensions and acousto-mechanical properties mimic an iliofemoral vein. The channel was perfused with plasma and the lytic&#160;recombinant tissue-type plasminogen activator. Bubble clouds were generated in the clot with a focused ultrasound source designed for the treatment of femoral venous clots. Motorized positioners were used to translate the source focus along the clot length. At each insonation location, acoustic emissions from the bubble cloud were passively recorded, and beamformed to generate passive cavitation images. Metrics to gauge treatment efficacy included clot mass loss (overall treatment efficacy), and the concentrations of D-dimer (fibrinolysis) and hemoglobin (hemolysis) in the perfusate. There are limitations to this in vitro design, including lack of means to assess in vivo side effects or dynamic changes in flow rate as the clot lyses. Overall, the setup provides an effective method to assess the efficacy of histotripsy-based strategies to treat DVT.

Histotripsy-aided lytic delivery or lysotripsy is under development for the treatment of deep vein thrombosis. An in vitro procedure is presented here to assess the efficacy of this combination therapy. Key protocols for the clot model, image guidance, and assessment of treatment efficacy are discussed.