Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove


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

doi: 10.3791/62133 Published: June 4, 2021
Aarushi Bhargava1, Samuel A. Hendley2, Kenneth B. Bader1,2


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 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.


Thrombosis is the condition of clot formation in an otherwise healthy blood vessel that obstructs circulation1,2. Venous thromboembolism has an annual healthcare cost of $7-10 billion, with 375,000-425,000 cases in the United States3. Pulmonary embolism is the obstruction of the pulmonary artery and is the most serious consequence of venous thromboembolism. The primary source of pulmonary obstruction is deep vein thrombi, primarily from iliofemoral venous segments4,5,6. Deep vein thrombosis (DVT) has inherent sequela besides pulmonary obstructions, with long term complications that result in pain, swelling, leg ulcerations, and limb amputations7,8,9. For critical obstructions, catheter directed thrombolytics (CDT) are the frontline approach for vessel recanalization10. The outcome of CDT depends on a number of factors, including thrombus age, location, size, composition, etiology, and patient risk category11. Moreover, CDT is associated with vascular damage, infections, bleeding complications, and long treatment time10. Next generation devices aim to combine mechanical thrombectomy with thrombolytics (i.e., pharmacomechanical thrombectomy)12,13. Use of these devices lower the lytic dosage leading to reduced bleeding complications, and shortened treatment time12,13,14 as compared to CDT. These devices still retain issues of hemorrhagic side-effects and incomplete removal of chronic thrombi15. An adjuvant strategy is thus needed that can remove the thrombus completely with lower bleeding complications.

One potential approach is histotripsy-aided thrombolytic treatment, referred to as lysotripsy. Histotripsy is a non-invasive treatment modality that uses focused ultrasound to nucleate bubble clouds in tissues16. Bubble activity is generated not via exogenous nuclei, but by the application of ultrasound pulses with sufficient tension to activate nuclei intrinsic to tissues, including clot17,18. The mechanical oscillation of the bubble cloud imparts strain to the clot, disintegrating the structure into acellular debris19. Histotripsy bubble activity provides effective degradation of retracted and unretracted blood clots both in vivo and in vitro20,21,22. Prior studies have23,24 demonstrated that the combination of histotripsy and the lytic recombinant tissue-type plasminogen activator (rt-PA) significantly increases treatment efficacy compared to lytic alone or histotripsy alone. It is hypothesized that two primary mechanisms associated with histotripsy bubble activity are responsible for the improved treatment efficacy: 1) increased fibrinolysis due to enhanced lytic delivery, and 2) hemolysis of red blood cells within the clot. The bulk of the clot mass is comprised of red blood cells24, and, therefore, tracking erythrocyte degradation is a good surrogate for ablation of the sample. Other formed clot elements are also likely disintegrated under histotripsy bubble activity but are not considered in this protocol.

Here, a benchtop approach to treat DVT in vitro with lysotripsy is outlined. The protocol describes critical operating parameters of the histotripsy source, assessment of treatment efficacy, and image guidance. The protocol includes designing a flow channel to mimic an iliofemoral venous segment and manufacturing human whole blood clots. The experimental procedure outlines the positioning of the histotripsy source and imaging array to achieve histotripsy exposure along the clot placed in the flow channel. Relevant insonation parameters to attain clot disruption and minimize off-target bubble activity are defined. The use of ultrasound imaging for guidance and assessment of bubble activity is illustrated24. Metrics to quantify treatment efficacy such as clot mass loss, D-dimer (fibrinolysis), and hemoglobin (hemolysis) are outlined23,24,25,26,27. Overall, the study provides an effective means for executing and assessing the efficacy of lysotripsy to treat DVT.

Subscription Required. Please recommend JoVE to your librarian.


For the results presented here, venous human blood was drawn to form clots after approval from the local internal review board (IRB #19-1300) and written informed consent provided by volunteer donors24. This section outlines a design protocol to assess lysotripsy efficacy. The protocol is based on a previous work by Bollen et al.24.

1. Clot modeling

NOTE: Prepare the clots within 2 weeks but more than 3 days prior to the day of the experiment to ensure clot stability and maximize retraction28. Prepare the clot following the approval from local institutional review board.

  1. Prepare borosilicate Pasteur pipette for storing the blood (see Table of Materials for specifications of the pipette). Borosilicate tubes are used because of the hydrophilic nature of the material which promotes platelet activation and clot retraction29. Seal the tip of the pipette via heating over a Bunsen burner.
  2. Draw fresh human venous blood. Aliquot the total blood drawn in 2 mL increments per desired clot. Transfer each 2 mL aliquot to one Pasteur pipette.
    NOTE: Execute step 1.2 within approximately 3 min of blood draw so that blood does not clot before transferring to pipettes. Also, ensure the volunteer donor is not on any medications that may alter the clotting cascade (e.g. blood thinners or platelet inhibitors).
  3. Incubate the blood aliquots within the pipettes (equal to the number of clots required) in a water bath for 3 h at 37 °C.
  4. Store the pipettes for a minimum of 3 days at 4 °C to allow for retraction of clots28. As the clots retract, serum will be observed to accumulate at the top of the clot within the pipette. The rt-PA response of clots remains stable for 2 weeks following retraction28.

2. Water tank preparation

  1. Fill the water tank with reverse osmosis water. Line the bottom surface of the tank with an acoustic absorbing material to reduce reflection of the therapy or imaging ultrasound pulses. Use a water handling system to degas and filter the water to minimize bubble nuclei.
    NOTE: One way to filter the water is using an inline filter. The specifications of the bag used for generating the representative results is given in the Table of Materials.
  2. Place two heating elements at the bottom surface of the tank. Heat the water to 37.3 ˚C to achieve the maximum lytic enzymatic activity30.
  3. Setup a flow channel as shown in Figure 1A. The flow channel consists of tubing, a model vessel with material and geometric properties representative of an iliofemoral vein, a reservoir for plasma, and a syringe on the distal most end of the reservoir (Table of Materials). The syringe is connected to a pump to regulate a flow through the channel during the experiment.
  4. Manually submerge the flow channel in the water tank to bring the channel to physiologic temperature during the degassing/filtering/heating stage (steps 2.1 and 2.2).

3. Preparation of plasma and rt-PA mixture

  1. Prior to experiment day
    ​NOTE: When stored at -80 °C, plasma is stable for at least 2.5 years31 and rt-PA is stable for at least 7 years32. Therefore, execute step 3.1 anytime within these periods to ensure stability of the two components.
    1. Dilute the rt-PA obtained from a manufacturer in powdered form to 1 mg/mL in sterile water.
    2. Aliquot 100 µL of diluted rt-PA into 0.5 mL centrifuge tubes and store them at -80 °C.
    3. Aliquot 35 mL of human fresh-frozen type O plasma in 50 mL centrifuge tubes. Store the tubes at -80 °C.
  2. On the experiment day
    1. Retrieve plasma aliquots from the freezer. Retrieve as many aliquots as the number of clots to be tested that day. Immerse the frozen aliquots in a water bath at 37 °C to thaw (~10 min).
    2. Once plasma has thawed, pour it into a beaker that is triply rinsed with ultrapure water. Lightly cover the mouth of the beaker with aluminum foil to prevent contamination and place the beaker in the water bath. Allow the foil to be loose enough to allow air to contact the plasma.
    3. Let plasma equilibrate at 37 °C to atmospheric pressure for at least 2 h.
    4. Take out the frozen rt-PA vials and place on ice until needed, with one vial for each experiment run.
    5. Make low gelling agarose (2%) in a 50 mL flask, by dissolving agarose in ultrapure water. Choose the total amount of agarose solution such that approximately 2 mL is available for each specimen to be analyzed. Heat the solution in flask in a microwave until bubbly. Secure the flask with a waterproof screw lid on it. Submerge the flask in the water bath alongside the plasma.
      ​NOTE: This step ensures agarose is available to secure exposed clot segments for histology analysis following histotripsy insonation.

4. Setting up histotripsy source and imaging array

  1. Ensure the motorized positioners can be controlled from the runtime environment of a programming platform, using directions and commands provided by the manufacturer. Check that the motors of the system are connected to the appropriate port of the computer with the runtime environment.
  2. Mount the histotripsy source on the motorized positioning system as shown in Figure 1B.
  3. Connect the histotripsy source to its driving electronics (e.g., power amplifier and function generator) via the appropriate connectors (e.g., BNC cables) as specified by the manufacturer.
    NOTE: Ensure that the driving electronics of the histotripsy source can be controlled via the runtime environment used in step 4.1.
  4. Cover the imaging array with a probe cover and affix the array coaxially in the aperture of the histotripsy source as shown in Figure 2. Be sure to understand the orientation of the imaging plane relative to the orientation of the histotripsy source.
  5. Connect the imaging array to an ultrasound scanning system. Ensure that this system can control the operation and triggering of the imaging array, and collect imaging data, as per the commands provided by the manufacturer of the scanner.
  6. Submerge the histotripsy source/imaging array into the tank while degassing as shown in Figure 1A. Gently remove any air bubbles using a syringe from the surface of the histotripsy source or imaging array.
    NOTE: Immerse the histotripsy source and imaging array completely in water before operation. Avoid touching the surface of the histotripsy source.
  7. Acquire B-mode images at a rate of 20 frames per second using the imaging array and the built-in commands of the scanner. Adjust the imaging window to ensure visualization of the focus of the histotripsy source in these real-time images.
    NOTE: It is assumed that the dimensions of the focal region of the therapeutic source are known.
  8. Set the operating parameters of the histotripsy source, including fundamental frequency (e.g., 1.5 MHz), pulse repetition frequency (e.g., 20-100 Hz), pulse duration (e.g., 1-20 cycles per pulse), and total number of pulses per location (e.g., 100-2,000)18,23,24,33. Modify these parameters if sufficient clot lysis is not achieved or if the bubble activity extends beyond the lumen of the model vessel. To set these parameters, use the protocol provided by the manufacturer of the source or use a programming platform that can communicate with the source (Step 4.3).
  9. Using the manufacturer's protocol or programming platform used in step 4.8, run the histotripsy source at the set parameters in degassed water only, without any obstruction in the surrounding environment. Increase the voltage applied to the histotripsy source until a bubble cloud is formed.
  10. Using the real-time imaging of step 4.7, adjust the position of the imaging array inside the confocal transducer opening until the bubble cloud is located approximately at the center of the image window. The bubble cloud is the region of hyperechoic pixels in the imaging plane (Figure 3). Tighten the screws to hold the imaging array firmly in the transducer opening.
    NOTE: If the array is aligned properly, the azimuthal position of the bubble cloud should be approximately at 0 mm in the imaging plane. The imaging array may project slightly from the inner surface of the therapy source, and therefore the range position of the bubble cloud may differ from the focal length of the source.
  11. Identify the bubble cloud location in the imaging plane. Assign the focus of the histotripsy source as the center of the bubble cloud (Figure 3).
  12. Record the detected focal location (step 4.11) in the imaging window (Figure 3). A possible way to mark the focal position is placing a cursor to note the location in the imaging window, if available with the imaging platform.
  13. Discontinue insonation and set the voltage applied to the histotripsy source to 0 V.

5. Clot preparation

  1. Remove the clot from the pipette by cutting the sealed end with pliers. Let the clot slide into a Petri dish along with the serum. If the clot does not dislodge, gently apply pressure from the other end of the pipette via saline flush to remove the clot.
  2. Cut the clot to 1 cm length using a scalpel, aiming for a uniform piece from the center (i.e., away from sections of the clot formed at the top or bottom of the pipette).
  3. Use a cleaning wipe to blot the cut section of the clot gently to remove excess fluid.
  4. Using tweezers, place the clot section gently on a weighing scale and record the weight.
  5. Manually raise the flow channel out of the water tank and remove the model vessel from the flow channel.
  6. Place the clot in the model vessel using tweezers and attach the model vessel again to the flow channel.
    NOTE: A nylon rod can be placed within the model vessel to prevent the clot from moving downstream due to the flow.
  7. Lower the flow channel into the tank in such a way that the proximal end of the stage relative to the reservoir is low compared to the distal side. The angling of the stage in this manner prevents trapping of bubbles in the model vessel when plasma is drawn through the flow channel in step 6.1.
  8. Add 30 mL of plasma to the reservoir using a pipette and monitor the temperature until it reaches at least 36 °C.
  9. Use a pipette to dispense the rt-PA (80.4 µg in 30 mL of plasma, 2.68 µg/mL) into the plasma reservoir. Stir the plasma with the pipette to ensure a uniform rt-PA distribution within the reservoir.

6. Priming the flow channel

  1. Draw plasma into the flow channel from the reservoir via the syringe pump until the plasma fills the model vessel.
    NOTE: If the clot is not flush with the nylon rod, use short pump draws at 60 mL/min to try to force the plasma downstream the clot or manually draw through the syringe. Limit the amount of plasma drawn in this process or refill the reservoir using additional plasma/rt-PA to ensure 30 mL of solution in the flow channel.
  2. Using the motorized positioners, align the imaging array parallel to the length of the clot using the imaging script (step 4.7). The parallel alignment enables the user to visually ensure proper placement of the clot and absence of bubbles inside the model vessel.
  3. Level the model vessel manually and visually ensure that no air bubbles are present using the imaging window (step 4.7).

7. Experiment procedure

  1. Pre-treatment
    ​NOTE: This step is to plan a path for the histotripsy source/imaging array for uniform histotripsy exposure along clot length.
    1. Align the imaging array using the motorized positioners such that the imaging plane is parallel to the cross-section of the clot (i.e., perpendicular to the orientation described in step 6.2).
    2. Under guidance via the imaging window (step 4.7), move the histotripsy source to the proximal end of the clot relative to the reservoir using the motorized positioners. At this point, adjust the histotripsy source position such that the marked focal point in step 4.12 aligns with the center of the clot.
    3. Determine the insonation path along the clot length. To define this path, set three waypoints along the length of the clot (i.e., positions of the motors where histotripsy bubble activity is contained within the clot) in 5 mm increments. Align the waypoints such that the overall motion of the histotripsy source along the path is antegrade with flow in the system (i.e., the first waypoint is at the most proximal end of the clot relative to the reservoir, and the third waypoint is in the distal position relative to the reservoir).
    4. Prior to finalizing each waypoint, fire test pulses from the histotripsy source with the same insonation parameters as step 4.8 but reduce the overall exposure to 10 total pulses. Adjust the position of the histotripsy source using the motorized positioners if necessary, to contain bubble activity within the clot.
    5. At each waypoint, save the motor positions using the commands provided by the manufacturer, similar to the step 4.1.
  2. Treatment
    ​NOTE: This step defines the procedure to treat the clot along its length according to the path defined in the pre-treatment step.
    1. Run the syringe pump at 0.65 mL/min and wait for meniscus of the plasma to move. This flow rate mimics a near total occlusion of the iliofemoral vasculature24,34.
    2. Interpolate the path created in step 7.1.3 with intermediate steps between the established waypoints with a fixed step size (e.g., 0.5 mm). The step size is chosen to be smaller than half the width of the focal region as measured along the clot length (elevational direction of the imaging array). Move the histotripsy source using motorized positioners at each path location with insonation parameters defined in step 4.8.
    3. Monitor/image bubble activity during application of the histotripsy pulse at each path location using the imaging window (step 4.7). Center the image on the histotripsy focus with the image dimensions covering 15 mm in the azimuth and range. Prior to the application of histotripsy pulse at each location, acquire a B-mode image to provide visualization of the clot and model vessel, by creating a script in a programming platform. Ensure the script establishes communication with the scanner using the manufacturer's commands.
    4. During the application of the histotripsy pulse, implement the acquisition of acoustic emissions in the script in step 7.2.3 to form passive cavitation images post hoc35. Acquire one passive cavitation image after every 10 treatment pulses. Apply a flat time gain compensation of 50 at 8 incremental depths till the end of the imaging depth. Choose an appropriate acquisition window size such that the entire signal from the clot is captured with minimum loss of energy due to windowing35.
    5. If off-target bubble clouds are present, adjust the transducer position in situ with the motorized positioners.
      ​NOTE: Monitor for missed triggers of the imaging array. Adjust the number of acquired imaging data sets to ensure the storage of data is completed before subsequent triggering.

8. Post experiment procedure

  1. Manually raise the model vessel out of the water tank to drain the perfusate via gravity. Be sure to keep the flow channel levelled to prevent the clot from moving downstream and out of the model vessel during draining.
  2. Collect the entire perfusate for further analysis in a small beaker (Figure 4A) by drawing plasma solution from the flow channel through the syringe pump at a very low flow rate.
  3. Disconnect the model vessel and remove the clot. If necessary, inject a small amount of saline into the model vessel to gently dislodge the clot.
  4. Wipe the clot similar to step 1.2.3. Weigh the clot on the weighing scale for assessing the clot mass loss.
  5. To analyze the D-dimer content, add 100 mg of aminocaproic acid to a microcentrifuge tube followed by 1 mL of perfusate, and mix well using a pipette. Perform a latex immune-turbidimetric assay to quantify the D-dimer within the sample36.
  6. To assess hemolysis, add 1 mL of perfusate to centrifuge tubes and spin at 610 x g (3,500 rpm) for 10 min. Combine 0.5 mL of supernatant (concentrate) with 0.5 mL of Drabkins solution and let the mixture rest at room temperature for 15 min. Transfer 200 µL to well plates, as shown in Figure 4B. Use a plate reader to read absorbance at 540 nm, Figure 4C (Drabkins assay37).
  7. Histology assessment
    1. Cut a section from the center of the clot of about 2-3 mm in length with a scalpel.
    2. Add the section to a cassette. Maintain orientation of the section relative to the direction of histotripsy pulse propagation.
    3. Add 2 mL of low gelling agarose solution prepared in step 3.2.5 into the cassette to fix the clot in place.
    4. Fix the sample in 10% formalin for 24 h. Place the sample in 70% alcohol after 24 h and perform standard hemotoxylin-eosin staining38.

9. Passive cavitation image analysis

  1. Process the signals acquired from the imaging array during the histotripsy excitation (step 7.2.4) using an algorithm based on the robust Capon beamformer39 to create an image of acoustic emissions generated by the bubble cloud at each treatment location.
    NOTE: To generate quantitative images, follow the steps described in Haworth et al.35. Otherwise, each pixel value in the image should represent the relative bubble cloud acoustic energy (units of V2) at each corresponding location.
  2. Segment the B-mode image aquired in steps 7.2.3 to distinguish between the pixels representing clot and the model vessel.
  3. Co-register the passive cavitation image with the B-mode image as shown in Figure 5B. Sum up the acoustic energy within the clot over the exposure period35.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

The protocol outlined in this study highlights the details of venous clot modeling, lysotripsy for clot disruption, and ultrasound imaging in an in vitro setup of DVT. The adopted procedure demonstrates the steps necessary to assess clot disruption due to the combined effects of rt-PA and histotripsy bubble cloud activity. The benchtop setup was designed to mimic the characteristics of a venous iliofemoral vein. Figure 1A shows a model vessel that has the acoustic, mechanical, and geometrical properties of the iliofemoral vein. The clot is placed inside the model vessel to mimic a partially occlusive thrombus. The clot is perfused with plasma and rt-PA drawn from a reservoir at a rate of 0.65 mL/min. This rate is consistent with slow flow rate in a highly occluded vessel34.

An elliptically focused transducer of 1.5 MHz fundamental frequency with a 9 cm major axis, 7 cm minor axis, and 6 cm focal length (Figure 2A) is mounted on the positioning system as noted in Figure 1B. An imaging array covered with ultrasound gel and a latex cover (Figures 2B,C) is mounted coaxially with the transducer as shown in Figure 1A via an opening in the center of the histotripsy source. The motorized positioners were used to translate the therapy transducer/imaging array along the clot length within the model vessel (Figure 1). Upon application of sufficient voltage to the histotripsy source, a bubble cloud is generated in the focal region of the transducer and visualized via ultrasound imaging, as shown in Figure 3. The focal position is defined as the center of the bubble cloud using the imaging plane (steps 4.10-4.11).

Figure 4A shows perfusates collected for two different treatment conditions. The beaker labeled as control contains perfusate of a clot exposed to plasma alone. The second beaker labeled as treated contains the perfusate of the lysotripsy treated clot. The collected perfusates are used to assess the hemoglobin (metric of hemolysis) and D-dimer (metric of fibrinolysis) content through assays as specified in the protocol. The difference in color of the perfusates denotes variability in hemoglobin concentration, which can be quantified via optical absorbance. The relationship between absorbance value and hemoglobin concentration can be determined through a calibration curve. Solutions with known hemoglobin content ranging from 0 (blank measurement) to 180 mg/mL are placed in the well plate and absorbance is determined in triplicate using the plate reader (Figure 4B,C). The upper absorbance limit of the plate reader may vary and may not be known a priori to making the solutions in the well plate. As such, hemoglobin concentrations up to 180 mg/mL are made in the well plate, Figure 4B. However, the plate reader used here can read absorbance for concentrations up to 23 mg/mL only, Figure 4C.

Figure 5A shows visualization of the clot within the model vessel via B-mode imaging prior to histotripsy exposure as specified in step 7.2.3. This image is acquired to determine the clot position for segmentation of the passive cavitation image. Figure 5B shows the passive cavitation image co-registered with the B-mode image acquired prior to histotripsy exposure. This figure confirms that acoustic energy is contained primarily within the clot during histotripsy exposure.

Typical clot disruption due to histotripsy and lytic are indicated in Figure 6. Figure 6A,B show the untreated and lysotripsy treated clot images, respectively. For samples exposed to histotripsy, disruption is primarily restricted to the clot center, consistent with the observed locations of bubble activity tracked with passive cavitation imaging (Figure 5B). However, with addition of lytic, mass loss also occurs in regions closer to the periphery of the clot. It is hypothesized that this additional mass loss is due to enhanced fluid mixing of the lytic under bubble activity. Fluid mixing increases the distribution and penetration depth of the lytic into the clot. Since the lytic is responsible for fibrinolysis40, the mass loss increases. Fibrinolysis can be quantified by measuring the D-dimer content in the perfusate41.

Figure 1
Figure 1: Experimental setup for lysotripsy of human blood clot. (A) The components of the setup are (1) focused histotripsy source with elliptical geometry, (2) latex-covered imaging array, (3) model vessel attached to flow channel, (4) flow channel, (5) reservoir, (6) acoustic absorbing material, (7) heating element, and (8) water tank filled with degassed and heated reverse osmosis water. The azimuth dimension of the imaging plane is perpendicular to the elevational and range dimensions (into the page). (B) The histotripsy source mounted on the motorized positioning system. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Ultrasound source and imaging components. Individual zoomed images of (A) focused histotripsy source, (B) imaging array, and (C) imaging array with ultrasound gel and latex cover. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Histotripsy bubble cloud visualized using imaging array. A bubble cloud is generated in the focal zone of the histotripsy source and imaged using an imaging array. The designated focus, shown as a cross, is saved for treatment planning. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Quantification of hemoglobin released due to clot lysis. (A) Perfusate samples collected following control study with plasma alone (no histotripsy or lytic), and treatment arm, histotripsy (e.g., 35 MPa peak negative pressure, 5 cycle pulse duration, 1.5 MHz fundamental frequency), and 2.68 µg/mL lytic exposure. (B) Well plate containing dilutions of known hemoglobin concentrations ranging from 180 mg/mL (top row, left-most corner) to 0 mg/mL (bottom row, right-most corner). The arrowhead points toward decreasing hemoglobin concentration. (C) These samples are used to create a standard curve to quantify hemoglobin produced due to histotripsy exposure via spectrophotometry. Absorbance curve for hemoglobin concentrations ranging from 0 to 23 mg/mL is obtained due to limitation of the plate reader in analyzing higher concentrations. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Images of the clot during treatment. (A) B-mode image acquired before the start of treatment pulse showing the clot position within the model vessel. (B) Post-hoc visualization ofacoustic energy emission calculated from passive cavitation imaging shown in hot colormap co-registered with B-mode image of the clot acquired prior to application of the histotripsy pulse. Please click here to view a larger version of this figure.

Figure 6
Figure 6: Histology of the ablated clot under different treatment conditions. (A) Control clot without treatment. (B) Clot treated with lysotripsy (e.g., 35 MPa peak negative pressure, single cycle pulse duration, 1.5 MHz fundamental frequency). The histotripsy pulse propagated from top to bottom in this image. The path for the histotripsy source along the length of the clot (i.e., perpendicular to the plane of the image shown here) is defined in step 7.2.3. The scale of the micrographs is 2 mm. Note that the degree of clot disruption achieved here would be reduced compared to insonation schemes with longer pulse duration24Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.


The proposed protocol presents a model to quantify treatment efficacy of lysotripsy. While the key details have been discussed, there are certain critical aspects to consider for the success of this protocol. The enzymatic activity of rt-PA has an Arrhenius temperature dependence30. Temperature is also a contributing factor to the speed of sound in water and tissue, and variations in temperature can cause minor alterations of the focal zone geometry. Thus, the water temperature should be carefully regulated at 37 ˚C. The dose of rt-PA used in the protocol (2.68 µg/mL) is consistent with that employed clinically for other pharmacomechanical thrombectomy strategies42. In step 5.8, 30 mL of plasma is transferred to the reservoir whereas a 35 mL aliquot is noted in step 3.1.3. This additional plasma accounts for loss in plasma due to evaporation over the course of hours when warmed to 37 ˚C for equilibration to atmospheric pressure.

The focal length, aperture width, and frequency of the therapy transducer dictate the size and depth of the focal region. Therefore, the transducer should be chosen such that these characteristics align with the diameter and the depth of the target vessel (e.g. femoral vein: 2-4 cm in depth and 0.6-1.2 cm in diameter)43. The extent of mechanical ablation is restricted to the extent of the bubble cloud. Thus, an understanding of the role insonation parameters play in modifying histotripsy bubble cloud behavior is critical33,44,45. The frequency and the strength of acoustic field should also be chosen noting the magnitude of attenuation due to medium and intermediary materials (e.g., model vessel). To ensure confinement of bubble activity with the target vessel, an appropriate imaging window should be chosen to monitor the focal zone. The operating parameters of the transducer should be chosen to avoid off target effects while maximizing mechanical clot disruption. In this protocol, mass loss was considered a primary metric of treatment efficacy. Increases in mass loss have been observed as the peak negative pressure or the duration of the histotripsy pulse are increased24,46, with a maximum observed mass loss of 94%. The presence of residual clot for investigated treatment arms facilitates comparison of therapeutic efficacy. However, insonation schemes to ensure total removal of the thrombus can also be devised.

The acoustic impedance (approximately 1.58 MRayl47,48) and the geometrical properties (0.6-1.2 cm in diameter43) of the model vessel should be representative of the iliofemoral venous vasculature (see Table of Materials for details). Polydimethylsiloxane and polyurethane are some of the other materials suitable to model the venous system based on their acousto-mechanical properties. In step 7, it is important to remove all the air bubbles from the model vessel to avoid shielding the clot from histotripsy exposure. For a model vessel of hydrophobic material, bubble clouds may form preferentially near the vessel wall instead of the center of the clot. Therefore, continuous monitoring of the bubble cloud should be done during the treatment via ultrasound imaging, and the transducer should be repositioned if necessary. Pilot studies should be conducted to determine histotripsy insonation parameters (e.g., pulse duration and peak pressure) that achieve the final intended clot disruption.

The imaging array is used to capture B-mode images and passive cavitation images for treatment visualization and to quantify bubble activity. B-mode imaging allows visualization of the model vessel and the clot, and passive cavitation imaging gauges the energy of the bubble activity associated with clot ablation24,49. The bandwidth of the imaging array should align with the desired bubble cloud activity with a high signal-to-noise ratio. For obtaining purely broadband signals associated with the inertial collapse of bubbles within the cloud, the bandwidth of the array should not coincide with the fundamental frequency of the transducer50,51. Histotripsy pulses are highly nonlinear52, and it is likely that harmonics of the fundamental frequency will be present in the received signal. The imaging system should be programmed to trigger on based on the known time of flight of the histotripsy pulse from the source to the focal zone to ensure collection of complete passive cavitation imaging data throughout the insonation. These signals should then be processed post hoc as discussed in steps 7.2.3 and 9 of the protocol.

It should be noted that the amount of hemolysis is sensitive to the handling of the clot. Therefore, care should be taken to minimize damage to the clot before treatment. To ensure reproducibility, clot modelling (step 1) and pre-treatment time (steps 6 and 7.1) should be same for all the clots treated with or without histotripsy exposure. In the post-treatment step of hemolysis assessment, it should be noted that plasma has its own absorbance. Therefore, the diluent used to form standard curves (e.g., optical absorbance vs. hemoglobin) should be formed using the same fluid used as the perfusate in the flow channel (e.g., in this study, plasma was used as the diluent to form standard curves).

This protocol aims to provide a benchtop setup to gauge the efficacy of lysotripsy to treat human whole blood clots. There are certain limitations that arise due to the in vitro nature of the set up. The acute clots used for this protocol consisted mainly of red blood cells and fibrin, making the approach of lysotripsy effective for DVT. However, later stages of thrombus may develop a stiff collagenous network53 that may resist the lysotripsy treatment due to the fibrin-specific nature of rt-PA. When treating in vivo, the primary clinical endpoint for treatment efficacy is restoration of flow. Mass loss was a primary metric for treatment efficacy in the in vitro protocol described here. Although flow was not assessed in this protocol, color Doppler imaging can be additionally incorporated along with passive cavitation imaging in step 7.2.4 to monitor flow restoration. The setup in this protocol uses a fixed flow rate, mimicking the flow rate in a highly occluded vessel, during the entire treatment in step 7.2. In vivo, vascular flow will increase as the clot disintegrates during the treatment. The additional shear stresses associated with increased flow will accentuate the clot degradation profile54. In vivo off-target effects cannot be ascertained in this setup, such as bleeding due to systemic administration of lytic55, vessel wall damage or vasospasm due to bubble cloud activity22. The in vitro nature of this study also limits the ability to assess long-term outcomes, such as vessel patency or re-thrombosis after treatment. The administration of lytic in this study mimicked systemic thrombolytics, whereas catheter-directed lytics is the preferred intervention for venous thrombosis7,14. Tissue attenuation can affect the histotripsy field and the imaging quality for in vivo studies, whereas here the acoustic path is primarily through degassed water. Processing of cavitation emission data with the robust Capon beamformer (step 9 of the protocol) is computationally expensive and was conducted off-line for post hoc analysis. Other beamformers (e.g., delay-and-sum35 or angular spectrum56) can be operated alternatively to provide real-time feedback, albeit with reduced range resolution.

In summary, this protocol presents a non-invasive approach to achieve deep vein thrombolysis of human blood clots. The protocol establishes a convenient and easy-to-replicate procedure for modeling of blood clots, treating them with lysotripsy, and simultaneous imaging during treatment. The protocol steps specifying histotripsy bubble cloud generation, treatment planning, and image guidance can be further used to investigate in vitro treatments of breast tumor, pancreatic tumor, and benign prostatic hyperplasia, where histotripsy has been shown to be more effective as compared to standard procedures57,58. The use of rt-PA in this protocol can be generalized to other drugs or drug carriers that are used for treating such tumors, along with histotripsy to increase the lytic efficacy.

Subscription Required. Please recommend JoVE to your librarian.


The authors have nothing to disclose.


This work was funded by the National Institutes of Health, Grant R01HL13334. The authors would like to thank Dr. Kevin Haworth for assisting with Drabkin's assay and Dr. Viktor Bollen for his support in designing the protocol. The authors are also thankful to Dr. Adam Maxwell for his guidance on designing the histotripsy source.


Name Company Catalog Number Comments
Absorbing sheets Precision acoustics F28-SMALL-M 300mm x 300 mm x 10 mm
Borosilicate Pasteur pippettes Fisher Scientific 1367820A 14.6 cm length, 2 mL capacity
Centrifuge tubes Eppendorf 22364111 1.5 mL capacity
Drabkin's assay Sigma Aldrich D5941-6VL
Draw syringe Cole-Parmer EW-07945-43 60 mL capacity
Filter bags McMaster-Carr 5162K111 Remove particle size upto 1 microns
Flow channel tubing McMaster-Carr 5154K25 Polyethylene-lined EVA plastic tubing (Outer diameter: 3/8", Inner diameter: 1/4"
Heating elements Won Brothers HT 300 Titanium Titanium rods placed at the bottom of tank
Imaging array Verasonics L11-5v 128 element with sensitivity from -55 to -49 dB
Low gelling agarose Millipore Sigma A9414
Model vessel McMaster-Carr 5234K98 6.6 cm length, 0.6 cm inner diameter, 1 mm thickness
Nanopure water Barnstead Nanopure Diamond ASTM type I, 18 Mohm-cm resistivity
Plasma Vitalant 4PF000 Plasma frozen within 24 hours
Plate reader Biotek Synergy Neo HST Plate Reader For haemoglobin quantification
Probe cover Civco 610-362
Programming platform MATLAB (the Mathworks, Natick, MA, USA)
Recombinant tissue-type plasminogen activator (rt-PA) Genentech Activase
Reservoir Cole-Parmer EW-07945-43 60 mL capacity
Syringe pump Cole-Parmer EW-74900-20 pump attached to the syringe to draw the flow in the flow channel at a pre-determined fized rate
Transducer In-house customized Eight-element, elliptically-focused transducer (9 cm major axis, 7 cm minor axis and 6 cm focal length), powered by custom designed and built class D amplifier and matching network
Ultrasound scaning system Verasonics Vantage Research Ultrasound System
Water tank Advanced acrylics C133 14 x 14 x 12, 1/2"



  1. Oklu, R. Thrombosis. Cardiovascular Diagnosis and Therapy. 7, Suppl 3 131-133 (2017).
  2. Satoh, K., Satoh, T., Yaoita, N., Shimokawa, H. Recent advances in the understanding of thrombosis. Arteriosclerosis, Thrombosis, and Vascular Biology. 39, (6), 159-165 (2019).
  3. Grosse, S. D., Nelson, R. E., Nyarko, K. A., Richardson, L. C., Raskob, G. E. The economic burden of incident venous thromboembolism in the United States: A review of estimated attributable healthcare costs. Thrombosis Research. 137, 3-10 (2016).
  4. Hirsh, J., Hoak, J. Management of deep vein thrombosis and pulmonary embolism. Circulation. 93, (12), 2212-2245 (1996).
  5. Browse, N. L., Clemenson, G., Croft, D. N. Fibrinogen-detectable thrombosis in the legs and pulmonary embolism. British Medical Journal. 1, (5908), 603-604 (1974).
  6. Plate, G., Ohlin, P., Eklöf, B. Pulmonary embolism in acute iliofemoral venous thrombosis. British Journal of Surgery. 72, (11), 912-915 (1985).
  7. Chen, J. X., Sudheendra, D., Stavropoulos, S. W., Nadolski, G. J. Role of catheter-directed thrombolysis in management of iliofemoral deep venous thrombosis. Radiographics. 36, (5), 1565-1575 (2016).
  8. Kahn, S. R., Solymoss, S., Lamping, D. L., Abenhaim, L. Long-term outcomes after deep vein thrombosis: postphlebitic syndrome and quality of life. Journal of General Internal Medicine. 15, (6), 425-429 (2000).
  9. Oğuzkurt, L., Ozkan, U., Gülcan, O., Koca, N., Gür, S. Endovascular treatment of acute and subacute iliofemoral deep venous thrombosis by using manual aspiration thrombectomy: long-term results of 139 patients in a single center. Diagnostic and Interventional Radiology. 18, (4), 410-416 (2012).
  10. Lauw, M. N., Büller, H. R. Current Approaches to Deep Vein Thrombosis. 136-160 (2014).
  11. Kearon, C., et al. Antithrombotic therapy for VTE disease: antithrombotic therapy and prevention of thrombosis: American college of chest physicians evidence-based clinical practice guidelines. Chest. 141, (2), 419-496 (2012).
  12. Pouncey, A. L., et al. AngioJet Pharmacomechanical Thrombectomy and Catheter Directed Thrombolysis vs. Catheter Directed Thrombolysis Alone for the Treatment of Iliofemoral Deep Vein Thrombosis: A Single Centre Retrospective Cohort Study. European Journal of Vascular and Endovascular Surgery. (2020).
  13. Tang, T., Chen, L., Chen, J., Mei, T., Lu, Y. Pharmacomechanical thrombectomy versus catheter-directed thrombolysis for iliofemoral deep vein thrombosis: a meta-analysis of clinical trials. Clinical and Applied Thrombosis/Hemostasis. 25, (2019).
  14. Kuo, T. -T., Huang, C. -Y., Hsu, C. -P., Lee, C. -Y. Catheter-directed thrombolysis and pharmacomechanical thrombectomy improve midterm outcome in acute iliofemoral deep vein thrombosis. Journal of the Chinese Medical Association. 80, (2), 72-79 (2017).
  15. Donaldson, C. W., et al. Thrombectomy using suction filtration and veno-venous bypass: single center experience with a novel device. Catheterization and Cardiovascular Interventions. 86, (2), 81-87 (2015).
  16. Khokhlova, V. A., et al. Histotripsy methods in mechanical disintegration of tissue: Towards clinical applications. International Journal of Hyperthermia. 31, (2), 145-162 (2015).
  17. Bader, K. B., Vlaisavljevich, E., Maxwell, A. D. For whom the bubble grows: Physical principles of bubble nucleation and dynamics in histotripsy ultrasound therapy. Ultrasound in Medicine & Biology. 45, (5), 1056-1080 (2019).
  18. Maxwell, A. D., et al. Noninvasive thrombolysis using pulsed ultrasound cavitation therapy-histotripsy. Ultrasound in Medicine & Biology. 35, (12), 1982-1994 (2009).
  19. Xu, Z., et al. Size measurement of tissue debris particles generated from pulsed ultrasound cavitational therapy-histotripsy. Ultrasound in Medicine & Biology. 35, (2), 245-255 (2009).
  20. Vlaisavljevich, E., et al. Effects of tissue stiffness, ultrasound frequency, and pressure on histotripsy-induced cavitation bubble behavior. Physics in Medicine and Biology. 60, (6), 2271-2292 (2015).
  21. Zhang, X., et al. Histotripsy thrombolysis on retracted clots. Ultrasound in Medicine & Biology. 42, (8), 1903-1918 (2016).
  22. Maxwell, A. D., et al. Noninvasive treatment of deep venous thrombosis using pulsed ultrasound cavitation therapy (histotripsy) in a porcine model. Journal of Vascular and Interventional Radiology. 22, (3), 369-377 (2011).
  23. Bader, K. B., et al. Efficacy of histotripsy combined with rt-PA in vitro. Physics in Medicine and Biology. 61, (14), 5253-5274 (2016).
  24. Bollen, V., et al. In vitro thrombolytic efficacy of single- and five-cycle histotripsy pulses and rt-PA. Ultrasound in Medicine & Biology. 46, (2), 336-349 (2020).
  25. Wang, Y. N., Khokhlova, T., Bailey, M., Hwang, J. H., Khokhlova, V. Histological and biochemical analysis of mechanical and thermal bioeffects in boiling histotripsy lesions induced by high intensity focused ultrasound. Ultrasound in Medicine & Biology. 39, (3), 424-438 (2013).
  26. Weisel, J. W., Litvinov, R. I. Fibrin formation, structure and properties. Sub-Cellular Biochemistry. 82, 405-456 (2017).
  27. Devanagondi, R., et al. Hemodynamic and hematologic effects of histotripsy of free-flowing blood: implications for ultrasound-mediated thrombolysis. Journal of Vascular and Interventional Radiology: JVIR. 26, (10), 1559-1565 (2015).
  28. Holland, C. K., Vaidya, S. S., Datta, S., Coussios, C. -C., Shaw, G. J. Ultrasound-enhanced tissue plasminogen activator thrombolysis in an in vitro porcine clot model. Thrombosis Research. 121, (5), 663-673 (2008).
  29. Sutton, J. T., Ivancevich, N. M., Perrin, S. R., Vela, D. C., Holland, C. K. Clot retraction affects the extent of ultrasound-enhanced thrombolysis in an ex vivo porcine thrombosis model. Ultrasound in Medicine & Biology. 39, (5), 813-824 (2013).
  30. Shaw, G. J., Dhamija, A., Bavani, N., Wagner, K. R., Holland, C. K. Arrhenius temperature dependence of in vitro tissue plasminogen activator thrombolysis. Physics in Medicine & Biology. 52, (11), 2953 (2007).
  31. Pinto, J., et al. Human plasma stability during handling and storage: impact on NMR metabolomics. Analyst. 139, (5), 1168-1177 (2014).
  32. Shaw, G. J., Sperling, M., Meunier, J. M. Long-term stability of recombinant tissue plasminogen activator at -80 C. BMC Research Notes. 2, (1), 117 (2009).
  33. Maxwell, A. D., et al. Cavitation clouds created by shock scattering from bubbles during histotripsy. The Journal of the Acoustical Society of America. 130, (4), 1888-1898 (2011).
  34. Jensen, C. T., et al. Qualitative slow blood flow in lower extremity deep veins on doppler sonography: quantitative assessment and preliminary evaluation of correlation with subsequent deep venous thrombosis development in a tertiary care oncology center. Journal of Ultrasound in Medicine. 36, (9), 1867-1874 (2017).
  35. Haworth, K. J., Bader, K. B., Rich, K. T., Holland, C. K., Mast, T. D. Quantitative frequency-domain passive cavitation imaging. IEEE Transactions on Ultrasonics, Ferroelectrics, and Frequency Control. 64, (1), 177-191 (2017).
  36. Hamano, A., et al. Latex immunoturbidimetric assay for soluble fibrin complex. Clinical Chemistry. 51, (1), 183-188 (2005).
  37. Drabkin, D. L., Austin, J. H. Spectrophotometric studies II. Preparations from washed blood cells; nitric oxide hemoglobin and sulfhemoglobin. Journal of Biological Chemistry. 112, (1), 51-65 (1935).
  38. Fischer, A. H., Jacobson, K. A., Rose, J., Zeller, R. Hematoxylin and eosin staining of tissue and cell sections. CSH Protocols. 2008, (2008).
  39. Coviello, C., et al. Passive acoustic mapping utilizing optimal beamforming in ultrasound therapy monitoring. The Journal of the Acoustical Society of America. 137, (5), 2573-2585 (2015).
  40. Mori, K., Dwek, R. A., Downing, A. K., Opdenakker, G., Rudd, P. M. The activation of type 1 and type 2 plasminogen by type I and type II tissue plasminogen activator. Journal of Biological Chemistry. 270, (7), 3261-3267 (1995).
  41. Righini, M., Perrier, A., De Moerloose, P., Bounameaux, H. D-Dimer for venous thromboembolism diagnosis: 20 years later. Journal of Thrombosis and Haemostasis: JTH. 6, (7), 1059-1071 (2008).
  42. Hilleman, D. E., Razavi, M. K. Clinical and economic evaluation of the Trellis-8 infusion catheter for deep vein thrombosis. Journal of Vascular and Interventional Radiology: JVIR. 19, (3), 377-383 (2008).
  43. De Sensi, F., et al. Predictors of successful ultrasound guided femoral vein cannulation in electrophysiological procedures. Journal of Atrial Fibrillation. 11, (3), 2083 (2018).
  44. Vlaisavljevich, E., et al. Effects of ultrasound frequency and tissue stiffness on the histotripsy intrinsic threshold for cavitation. Ultrasound in Medicine & Biology. 41, (6), 1651-1667 (2015).
  45. Vlaisavljevich, E., et al. Histotripsy-induced cavitation cloud initiation thresholds in tissues of different mechanical properties. IEEE Transactions on Ultrasonics, Ferroelectrics, and Frequency Control. 61, (2), 341-352 (2014).
  46. Hendley, S. A., Paul, J. D., Bader, K. B. Mechanistic investigation of clot degradation via the action of histotripsy and thrombolytic. Joint AAPM | COMP Virtual Meeting. The American Association of Physics in Medicine. (2020).
  47. Goss, S. A., Johnston, R. L., Dunn, F. Comprehensive compilation of empirical ultrasonic properties of mammalian tissues. The Journal of the Acoustical Society of America. 64, (2), 423-457 (1978).
  48. Duck, F. A. Physical Properties of Tissues. Duck, F. A. Academic Press. 137-165 (1990).
  49. Bader, K. B., Haworth, K. J., Maxwell, A. D., Holland, C. K. Post hoc analysis of passive cavitation imaging for classification of histotripsy-induced liquefaction in vitro. IEEE Transactions on Medical Imaging. 37, (1), 106-115 (2018).
  50. Crake, C., et al. Enhancement and passive acoustic mapping of cavitation from fluorescently tagged magnetic resonance-visible magnetic microbubbles in vivo. Ultrasound in Medicine & Biology. 42, (12), 3022-3036 (2016).
  51. Gyongy, M., Coussios, C. Passive spatial mapping of inertial cavitation during HIFU exposure. IEEE Transactions on Biomedical Engineering. 57, (1), 48-56 (2010).
  52. Canney, M. S., Bailey, M. R., Crum, L. A., Khokhlova, V. A., Sapozhnikov, O. A. Acoustic characterization of high intensity focused ultrasound fields: A combined measurement and modeling approach. The Journal of the Acoustical Society of America. 124, (4), 2406-2420 (2008).
  53. Czaplicki, C., et al. Can thrombus age guide thrombolytic therapy. Cardiovascular Diagnosis and Therapy. 7, Suppl 3 186-196 (2017).
  54. Bajd, F., Vidmar, J., Blinc, A., Sersa, I. Microscopic clot fragment evidence of biochemo-mechanical degradation effects in thrombolysis. Thrombosis Research. 126, (2), 137-143 (2010).
  55. Wang, C., et al. Efficacy and safety of low dose recombinant tissue-type plasminogen activator for the treatment of acute pulmonary thromboembolism: a randomized, multicenter, controlled trial. Chest. 137, (2), 254-262 (2010).
  56. Arvanitis, C. D., Crake, C., McDannold, N., Clement, G. T. Passive acoustic mapping with the angular spectrum method. IEEE Transactions on Medical Imaging. 36, (4), 983-993 (2017).
  57. Khokhlova, V. A., et al. Histotripsy methods in mechanical disintegration of tissue: towards clinical applications. International Journal of Hyperthermia: The Official Journal of European Society for Hyperthermic Oncology, North American Hyperthermia Group. 31, (2), 145-162 (2015).
  58. Roberts, W. W. Development and translation of histotripsy: current status and future directions. Current Opinion in Urology. 24, (1), 104-110 (2014).
This article has been published
Video Coming Soon

Cite this Article

Bhargava, A., Hendley, S. A., Bader, K. B. An In vitro System to Gauge the Thrombolytic Efficacy of Histotripsy and a Lytic Drug. J. Vis. Exp. (172), e62133, doi:10.3791/62133 (2021).More

Bhargava, A., Hendley, S. A., Bader, K. B. An In vitro System to Gauge the Thrombolytic Efficacy of Histotripsy and a Lytic Drug. J. Vis. Exp. (172), e62133, doi:10.3791/62133 (2021).

Copy Citation Download Citation Reprints and Permissions
View Video

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
simple hit counter