Microbubble Write Up

Assessing the stability of microbubbles over time

Katherine Kiang

Bioacoustics Research Lab, Electrical and Computer Engineering, University of Illinois Urbana

Champaign

Background

Microbubbles, which are small gas filled bubbles between one micrometer and one

millimeter in diameter, have many potential biomedical applications in the diagnosis and

treatments of diseases. More specifically, microbubbles can be used as ultrasonic contrast agents

in order to image of many areas of the body, such as the heart, prostate, or liver [1] [2]. The

pressure caused by the high frequency of ultrasound scanner cause the microbubbles to contract

and expand. This causes the bubbles to be more reflective and therefore easier to see.

Additionally, ultrasound scanners can tune into the “overtones” created by the resonance of the

microbubbles, allowing for extremely targeted and specific imaging [3]. For better use of

microbubbles in this context, it is of interest to understand how these microbubbles change over

time. To do this, microbubbles were made, and the size, microbubble number density, collapse

threshold, and attenuation were studied over a course of six weeks. The collapse threshold refers

to the pressure needed to burst the microbubbles while the attenuation refers to the reduced

strength of a transducer signal as microbubbles are added.

Procedure and methods

Production of microbubbles [4]

A large batch of albumin based one micron bubbles were made before the experiment

started. The dextrose stock solution was made with 15g dextrose [Fisher Scientific], 100mL

18mΩ water, 0.12 g Na2HPO4 [Sigma-Aldrich], 0.02g KH2PO4 [Sigma-Aldrich], and 0.1g

NaN3 [Sigma-Aldrich. The BSA stock solution was made with 5g BSA [Sigma-Aldrich], 100mL

18mΩ water, and 0.01g NaN3.

Each batch of microbubbles was fabricated using 12ml of 15% dextrose and 4ml of 5%

BSA in a 50ml centrifuge tube. N-decafluorobutane gas was then added to the tube for about six

seconds in order to purge the tube of the air. Immediately after the tube was vortexed for a

minute and transferred to a 40 mL ultracentrifuge tube. More n-decafluorobutane was added for

a three seconds and the tube was sonicated for 55 seconds at 90% power (450 Watts) (Figure 1).

A metal tube holder was used to hold the 40mL ultracentrifuge tube during sonication. Once the

tube was sonicated, 10 mL of DPBS was added to the tube and it was inverted to mix a few times

and let to sit 5-10 minutes on a rack.

Figure 1: Mock set-up of microbubble sonication technique

After fabrication, the bubbles had to be separated by size. The bubbles were brought up

into a 20 mL syringe. Air bubbles were removed and care was taken so as not to draw up any of

the foam that forms on top of the bubbles during sonication. Once the bubbles were in the

syringe the needle was taken off and the syringe capped and left to sit vertical in the fridge for

one to six hours.

More DPBS was added to the remaining bubbles in the ultracentrifuge tube and allowed

to sit for 5-10 minutes, after which they were put into another syringe by the same process

described above. The microbubbles were then left in the fridge for 1-6 hours. During this time, a

white band formed near the top of the syringe as the microbubbles separated by size. Size

separation was confirmed by suspending the white band in 5 mL of DPBS to view under the

microscope.

Once the bubbles finished rising, there were two parts of the bubbles left. The top white

band, and the lower liquid fraction. The lower liquid fraction was the 1 micron bubbles desired

(Figure 2). Therefore, that part was transferred to a new syringe to rise overnight in the same

manner as before. After the bubbles rose a second time and the lower fraction transferred into a

new syringe, they were ready for use. Seven 8mL syringes were used for each week of counting,

sizing, and DPCD. Three tubes of 30mL were used for three different attenuation experiments.

Figure 2: Rising Microbubbles [4]

Counting and Sizing The Microbubbles

Counting and sizing was done the day after the microbubbles were fabricated and then

every week thereafter for six weeks.

To count the microbubbles, a small amount of microbubbles were loaded into a

hemocytometer. Frist, a syringe of the previously prepared microbubbles was taken out and

gently rolled in someone’s hands to re-suspend the bubbles. Then, a few drops of the bubbles

were put into a coulter counter tube. If the microbubbles needed to be diluted (which only

happened the first week), another tube was prepared with a small about of DPBS. In a third tube,

a 1:9 ratio of microbubble to DPBS was mixed by using a pipette to draw the mixture up and

down. Once the microbubbles were prepared, a pipette was used to draw up about 100

microliters of the bubbles (either straight up or diluted). This was carefully loaded into one side

of the cleaned and prepared hemocytometer.

Once the hemocytometer was prepared a phase contrast microscope was used to image

the microbubbles inside the hemocytometer. For the counting images, 20x magnification was

used.

Images were taken of all 25 squares on the hemocytometer using the microscopes software. One

image was taken focused on the bubbles, another focused on the hemocytometer squares.

Counting was done by overlaying the images on Power Point (taking advantage of

transparency settings) and manually counting using a counter. The microbubble density equation

is as follows:

Number density = (N*D)/(Q)

N = number manually counted; D = dilution factor Q = number of squares

To size the microbubbles, 10 images were taken at random at 100x on the same

microscope. These images were focused solely on the microbubbles and converted for RGB to

Greyscale before saving. These images were analyzed in the Matlab software (DPCD suite

bubble size) written by Daniel King [5]. To do this the bubble sizing software was opened and

each image individually loaded (load new image button). Once the image was opened, the detect

circles function was able to be used. Manual size adjustment was also possible if needed. In the

Matlab workspace the x-axis would give the radius of the bubbles in microns and the count

would say how many bubbles were at each size. An average of the microbubble radius was taken

by adding up the amount of bubbles at each radius and multiplying it by the radius. This number

was then divided by the total amount of bubbles sized. Excel was used to keep track of the

microbubble size data.

Finding the collapse threshold of microbubbles

Assessing the collapse threshold of the microbubbles was done the day after the

microbubbles were fabricated and then every week thereafter for six weeks.

Finding the collapse threshold was done through a DPCD experiment designed by Dr.

Daniel King [5]. To set up this experiment, a RITEC RAM-5000 (machine to control the

pressure) and a 5MHz transducer were connected to a computer via an already inserted A/D card.

Pressure settings for the RITEC were calibrated weekly.

To connect the RITEC and transducer to the computer a number of materials were needed

– a small tank filled with degassed water (placed above a stir palate, a PCD detector, the

transducers, and cables. A PCD detector with a 5 MHz (transmit) transducer in the middle and a

receiver transducer on each side was placed inside the tank (Figure 3).

Cables were connected as follows:

5 MHz transducer & RF Burst No. 2 – L/R side of attenuation bar, respectively

R/L Receiver Transducers (2) - Receiver A No. 1 and Receiver B No 2, respectively (should be

diagonal)

Receiver A/B RF Monitor (2) – A/D card channel 1/3 (respectively)

Sync (back of RITEC) – Sync (A/D card)

Figure 3: Mock DPCD Setup

Once the setup was complete, the RITEC was powered on and 4Ch bubble snapshot was

opened on the computer.

The RITEC settings were as follows:

Receiver: Gain-22dB, High pass filter-1MHz, Low pass filter-20MHz

Triggers and Gates: Internal source, PRF – 10 Hz

GA-2: Frequencey-4.6 MHz, Number of cycles: 3

A directory was made and the 4Ch bubble snapshot settings were as follows (most default):

Sampling rate: 100 MHz, Channel Mode: Dual Mode, Voltage Range: 1V, No. of Points: 16k,

Trigger Edge: Rising, Trigger Level: 10, Coupling: DC, Impedance: 50 Ohms; Time between

repeats: 0

Baseline values were then taken. The RITEC was turned to a setting of 10 and using 4Ch

bubble snapshot, the transducer focus was found (look for a peak around 47 milliseconds) by

moving the PCD detector around the tank with the addition of the 50 micron wire with weight on

the end.

Another base value was needed with the detector positioned with its top just below the

highest point in the tanks water level. This was done using clamps. A magnetic stir bar was then

inserted into the tank and the stir plate turned onto a low setting (3-4). Once the PCD was

properly in place, 50 scans were taken with no RITEC output and bubbles to act as a control.

Once all the baselines were taken the actual DPCD experiment was able to be conducted.

To run the experiment, microbubbles (which were re-suspended using the same method

described in the sizing procedure) were added a few drops at a time and 500 snapshots for each

of ten different RITEC settings (4, 1, 20, 4, 8, 10, 6, 12, 17, 15 – see Table 1 for corresponding

pressure values) were taken. This process was repeated 10 times. Bubbles were added when

signals could not be seen. Over the course of the experiment, bubbles were added so as to make

sure signals were seen at smaller settings.

Table 1: Corresponding Pressure Setting For Each RITEC Setting

These results were analyzed in the same DPCD suite used to size the bubbles. This time

however, the Analyze/Auto classify software was used. Once this software was opened, the

dataset from the DPCD experiment was loaded using load dataset (frequency=4.6) and auto

classified. Care was taken to adhere to the correct naming convention to make sure the auto

classifier ran properly. After the data was classified the Plot Post excitation curve software was

used to plot the data in a graph which correlated the RITEC pressure settings to the percentage of

microbubbles that burst at that setting. The auto classified files were loaded into this software

as .mat files (put in 5MHz for transducer) and the pos-excitation curve was plotted. Because

there were few to no signals at many of the low (1,2,4) settings, the curves didn’t fit well every

week. To help analyze the curves, they were plotted with a -10 db minimum threshold. The area

of interest in analyzing this data was the 50% threshold, in other words the pressure at which

50% of the microbubbles collapsed.

Attenuation of Microbubbles

Attenuation was tested after weeks 0, 3, and 6. This test used a 7.5 db transducer and a

UT340 Pulser Receiver System.

To setup the attenuation experiment a stir-plate was placed inside the largest tank in the

daedal room. Next a smaller tank was placed on top of the stir plate and filled with degassed

water. Then a hydrophone and the transducer were placed into the smaller, water filled tank

(Figure 4a, 4b). The hydrophone was toward the back and connected to its amplifier, which is

connected to Ch1-PDA 14 in the UTEX. The transducer in front is connected to Pulse Rec in the

UTEX. Finally, Sync Out from the computer is connected to Sync on the UTEX.

Figure 4: a) Mock Attenuation Setup: Eye Level b) Mock Attenuation Setup: Overhead

Similar to calibration, the focus of the transducer must be found. This is initially done by

hand and then fine-tuned using the Daedel Position Menu to move the Daedel and PDA 14 to see

the transducer signal. The UTEX settings when finding the focus are: Voltage- 200 V ; Pulse

Width- 40 ns; Rep Rate- 200 Hz; Internal; PE/Gain – 0; PC/Gain – 0 Mode – Pitch Catch.

Once the focus was found baseline snapshots were taken with the transducer at the focus,

12 micrometers in front of the focus, and 12 micrometers in back of the focus. These snapshots

were taken with the same UTEX settings as before except with 100 V and with varying widths.

The widths tested were: 2 ns, 4 ns, 6 ns, 8 ns. The transducer was then kept at 12 micrometers in

front of the focus and a plastic U shaped holder wrapped in Ceram Wrap (carefully, without

double layering and using rubber bands) and with a small stir bar inside was placed between the

transducer and hydrophone. To place the holder, one person lowed the holder while the other

poured in water using a beaker. The stir plate was then turned on. Once the holder was in place,

control snapshots of the images were taken at all the UTEX settings (the differing widths).

To begin the experiment, 6 mL of microbubbles were added by putting an 18G needle on

the microbubble syringe. Once microbubbles were added and allowed to disperse for about a

minute, images of the signal were taken at each UTEX setting. This was done for 3 time points.

Then another 6 mL was added and the process repeated. In the end, there was a 5x concentration.

Analyzation of the attenuation data was done by comparing the height of the signals at

each width as the microbubble concentration increased.

Results

Figure 5:

Microbubble concentration for each week

Figure 6: The radius (in micrometers) of the microbubbles each week of testing

0 1 2 3 4 5 6 71.00E+06

1.00E+07

1.00E+08

1.00E+09

f(x) = − 37760714.2857143 x + 212705000R² = 0.754028853044046

Weekly microbubble concentration

CountLinear (Count)

Week

Mic

robu

bble

Con

cent

ratio

n

0 1 2 3 4 5 6 70

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

f(x) = 0.0273861437874725 x + 0.515900755273104R² = 0.910859135485639

Weekly Microbubble Size

SizeLinear (Size)

Week

Miro

bubb

le S

ize (R

adiu

s)

Figure 7: 50% Postecitation each week of testing

1 2 3

-1

-0.8

-0.6

-0.4

-0.2

0

0.2

0.4

0.6

0.8

Attenuation Over Time UTEX 2 Setting

Series1Linear (Series1)Series2Linear (Series2)Series3Linear (Series3)Series4Linear (Series4)Series5Linear (Series5)

Week

Atten

uatio

n Am

ount

Figure 8: Attenuation amount each week for UTEX 2 (Series # is Concentration)

0 1 2 3 4 5 6 70

0.51

1.52

2.53

3.54

4.5

f(x) = 0.142857142857143 x + 2.78571428571429R² = 0.25974025974026

Weekly 50% Posexcitation

50% PosexcitationLinear (50% Posexcitation)

Week (**week 5 N/A because didn't hit 50% postexcitation)

RITE

C Se

tting

With

50%

Pos

texc

itatio

n

1 2 3

-1.6-1.4-1.2

-1-0.8-0.6-0.4-0.2

00.20.4



Week

Atten

uatio

n Am

ount



1 2 3

-0.6-0.4-0.2

00.20.40.60.8

11.21.4



Week

Atten

uatio

n Am

ount

1 2 3

-1

-0.5

0

0.5

1

1.5

2



Week

Atten

uatio

n Am

ount


Analysis and Discussion

After running an ANOVA on the microbubble count data, a p value of 0.016039 was

found, suggesting a statistically significant difference in the microbubble concentration over

time. The microbubble size data had a p value of 0.012359, suggesting a statistically significant

difference in the microbubble size over time. This implies that the microbubbles were not stable

in their concentration and size over the duration of the experiment. A possible explanation for

this instability would be that the bubbles burst during storage. Since the average microbubble

size increased, it would indicate that the smaller microbubbles burst first.

Looking at the post excitation curve data over time, there doesn’t seem to be a statistical

significance, as the p value was 0.597098. This implies stability of the microbubbles over time in

regards to their collapse threshold. However, this could be related to not seeing many at low

RITEC settings during DPCD, which could lead to a less accurate post excitation curve.

When analyzing the attenuation data, the attenuation amounts had an inconsistent mixture

of negative and positive values for the same concentration and UTEX settings. A positive value

implies the signal shrunk and vice versa. Therefore, positive and negative numbers do not make

sense. What should be seen is increasingly positive numbers as the concentration of

microbubbles increased.

Conclusions

Overall, this experiment showed some significant differences in microbubble count and

concentration over time. This instability seems to contradict the hypothesis that the albumin

based microbubbles are stable over time. However, there seems to be stability in the collapse

threshold of the microbubbles over time. This inconsistency suggests the need more research to

be done, possibly with microbubbles at higher concentrations. Additionally, the attenuation

experiment must be assessed to see if there are problems with the experimental method causing

the illogical data.

Sources:

[1] "European Heart Journal - Cardiovascular Imaging." Microbubbles and Ultrasound: From

Diagnosis to Therapy. N.p., n.d. Web.

[2] F. J. Fry, N. T. Sanghvi, R. S. Foster, R. Bihrle and C. Hennige. Ultrasound and

Microbubbles: Their Generations, Detection and Potential Utilization in Tissue and Organ

Therapy - Experimental. Ultrasound in Medicine and Biology, 21 1227-1237, 1995.

[3] Blomley, Martin J K, Jennifer C. Cooke, Evan C. Unger, Mark J. Monaghan, and David O. Cosgrove. "Microbubble Contrast Agents: A New Era in Ultrasound." BMJ : British Medical Journal. BMJ, n.d. Web.

http://www.brl.uiuc.edu/Publications/1995/Fry-UMB-1227-1995.pdf



[4] Borrelli, Michael J., and William D. O'Brien. "Result Filters." National Center for Biotechnology Information. U.S. National Library of Medicine, n.d. Web.

[5] D. A. King and W. D. O’Brien, Jr., “Quantitative Analysis of Ultrasound Contrast Agent

Postexcitation Collapse,” IEEE Trans UFFC 61:7, 1237-1240 (2014).

Documents

Microbubble Write Up