Fabricating and Labeling Microbubbles with Fluorescent and Radioactive Tracers

Summary

This protocol outlines the fabrication of lipid microbubbles and a compatible one-pot microbubble radiolabeling method with purification-free >95% labeling efficiency that conserves microbubble physicochemical properties. This method is effective across diverse lipid microbubble formulations and can be tailored to generate radioactive and/or fluorescent microbubbles.

Abstract

Microbubbles are lipid-shelled, gas-filled particles that have evolved from vascular ultrasound contrast agents into revolutionary cancer therapy platforms. When combined with therapeutic focused ultrasound (FUS), they can safely and locally overcome physiological barriers (e.g., blood-brain barrier), deliver drugs to otherwise inaccessible cancers (e.g., glioblastoma and pancreatic cancer), and treat neurodegenerative diseases. The therapeutic arsenal of microbubble-FUS is advancing in new directions, including synergistic combination radiotherapy, multimodal imaging, and all-in-one drug loading and delivery from microbubble shells.

Labeling microbubbles with radiotracers is key to establishing these expanded theranostic capabilities. However, existing microbubble radiolabeling strategies rely on purification methodologies known to perturb microbubble physicochemical properties, use short-lived radioisotopes, and do not always yield stable chelation. Collectively, this creates ambiguity surrounding the accuracy of microbubble radioimaging and the efficiency of tumor radioisotope delivery.

This protocol describes a new one-pot, purification-free microbubble labeling methodology that preserves microbubble physicochemical properties while achieving >95% radioisotope chelation efficiency. It is versatile and can be applied successfully across custom and commercial microbubble formulations with differing acyl lipid chain length, charge, and chelator/probe (porphyrin, DTPA, DiI) composition. It can be adaptively applied during ground-up microbubble fabrication and to pre-made microbubble formulations with modular customizability of fluorescence and multimodal fluorescence/radioactive properties. Accordingly, this flexible method enables the production of tailored, traceable (radio, fluorescent, or radio/fluorescent active) multimodal microbubbles that are useful for advancing mechanistic, imaging, and therapeutic microbubble-FUS applications.

Introduction

Microbubbles are micron-sized supramolecular theranostic agents with a gas core stabilized by a protein, polymer, or, in most cases, a lipid shell (Figure 1A). When injected into the bloodstream, microbubbles maintain gas/liquid interfaces that are detectable by ultrasound for minutes-long timeframes prior to the dissolution of their gas cores^1,2. Consequently, the first clinical use of microbubbles was as real-time ultrasound imaging contrast agents³. The invention of therapeutic focused ultrasound (FUS) expanded microbubble clinical utilities. When stimulated by low-frequency FUS, microbubbles oscillate and generate targeted, tunable mechanical forces ranging from transient vascular permeabilization to focal tissue ablation^4,5. As a result, over the last 20 years, microbubble-FUS has been explored for blood-brain barrier (BBB) opening, tumor (e.g., pancreatic, brain, and liver metastatic cancer) drug and imaging probe delivery, neurodegenerative disease therapy and cancer ablation^{6,7,8,9,10,11}.

The theranostic arsenal of microbubbles continues to advance in new and exciting directions. Conventional microbubble-FUS delivery applications rely on the co-administration of therapeutic or imaging cargo alongside commercial microbubbles. There is growing interest in enhancing microbubble-FUS delivery capabilities by understanding microbubble shell/biological interactions, exploring custom-made non-commercial microbubble formulations, and generating all-in-one theranostic microbubbles with cargo loaded directly onto the microbubble shell^12,13,14. In fact, approximately 40% of lipid microbubble drug delivery studies make use of such shell-loaded microbubbles¹⁵. Beyond imaging and drug delivery, microbubble-FUS has also shown promise in enhancing cancer radiotherapy¹⁶, and activating antineoplastic effects of otherwise benign shell-loaded agents through sonodynamic therapy^17,18.

These conventional and expanded directions in microbubble cancer applications can be more strategically advanced by labeling microbubble shells with radioactive tracers. In the realm of all-in-one cargo-loaded microbubbles, such radiolabeling 1) facilitates gold-standard, quantitative assessment of the on and off-target biodistribution of these loaded microbubble shells, 2) derives pharmacokinetic structure-activity relationships that inform optimal selection of microbubble compositions to maximize on-target delivery, and 3) guides strategic and appropriate image-guided application and treatment planning (e.g., types of tissue targets, dosimetry, drug selection to mitigate off-target safety concerns, utility compared to conventional co-treatment paradigms) of all-in-one cargo-loaded systems^15,19. At a preclinical stage, such an understanding of microbubble shell fate can also illuminate broader microbubble-FUS mechanisms of action. For example, lipid transfer from microbubble shells to target cells has been shown to influence FUS-enabled sonoporation^12,20. Understanding and optimizing such transfer can thus inform preclinical and clinical microbubble-FUS therapies in which sonoporation is implicated (in vitro transfection, drug delivery, tumor ablation, radiation sensitization, and sonodynamic therapy^{20,21,22,23,24,25}). Dual ultrasound and radioimaging facilities would also enable FUS vessel opening and treatment monitoring (e.g., BBB opening kinetics) from a single agent rather than conventional dual agent designs²⁶. In the same vein, lipid microbubble radiolabeling could serve as an all-in-one single-agent microbubble-FUS/radiotherapy alternative to microbubble-FUS + radiopharmaceutical co-delivery platforms²⁷.

The fragility of microbubbles is an untrivial challenge to such labeling. All existing radiolabeling strategies are limited by purification methodologies known to perturb microbubble stability and size, while some also feature ineffective and unstable radiolabeling^{28,29,30,31,32}. Purification requirements also lead to lengthier protocols. Combined with the use of short-lived radioisotopes (e.g., ¹⁸F t_1/2 1.8 h,^28,29 99mTc t_1/2 6 h,^32 68Ga t_1/2 1 h³¹), this creates inefficiencies related to radioisotope decay and confines radioimaging and treatment planning timeframes. Collectively, these limitations risk the acquisition of shortened and unrepresentative radioimaging, inaccurate pharmacokinetic data, and inefficient tumor radioisotope delivery.

In this report, these limitations are overcome by leveraging the strong and stable metal chelation capabilities of porphyrin. Porphyrins are organic, heterocyclic macromolecules with a highly conjugated planar ring and a central coordination site that can accommodate a variety of metals. This includes longer-lived radioisotopes such as copper-64 (t_1/2 12.7 h), a radiopharmaceutical with positron emission tomography (PET), and γ-counting feasibilities³³. When conjugated to a lipid backbone, porphyrins can be readily incorporated into supramolecular structures and subsequently labeled with copper-64 with speed, high chelation efficiency, and serum stability, while maintaining the properties of the parent unlabeled particles^33,34. Furthermore, porphyrins are fluorescently active with modular self-quenching in nano and microparticles that is restored upon particle disruption; a complementary readout to PET and γ-counting that facilitates both bulk and microscopic shell fate analysis (Figure 1A)¹⁵.

By using porphyrin-lipid as a chelator, these properties were exploited to generate a new one-pot, purification-free microbubble radiolabeling methodology (Figure 1B,C) that overcomes limitations associated with existing microbubble radiolabeling methods. This protocol achieves >95% copper-64 chelation efficiency, does not require post-labeling purification, and preserves microbubble physicochemical properties. It can be integrated easily into the "ground-up" fabrication of lipid microbubbles prior to their activation (Figure 1B). It is versatile and can be applied successfully across custom and commercial microbubble formulations with differing acyl lipid chain length (C16 to C22), charge (neutral and anionic), and porphyrin-lipid compositions (1 mol%, 10 mol%, 30 mol%), generating microbubbles with both radio and fluorescence activity. Its adaptability can also extend beyond porphyrin. The one-pot protocol can be modified to use alternative commercially available chelators (e.g., diethylenetriamine pentaacetate (DTPA)-lipid) and fluorophores (e.g., DiI). It can also be modified to label pre-made microbubble formulations through a "spiking" approach. Accordingly, this method enables the production of tailored, traceable (radio, fluorescent, or dual radio/fluorescent active) microbubbles useful for advancing mechanistic, imaging and therapeutic microbubble-FUS applications. The protocol below outlines the fabrication of lipid microbubbles, application of the one-pot radiolabeling protocol, requisite radiolabeling and physicochemical property characterization, and potential modifications.

Figure 1: Microbubble fabrication and radiolabeling protocol. (A) Porphyrin-lipid, in the form of pyropheophorbide-a-lipid, serves as a multimodal chelator within this protocol. As a monomer chelated to copper-64 (i), it has PET and imaging capabilities. Its fluorescence is quenched in particle form (microbubbles (ii) and their post-dissolution nanoprogeny (iii)) and unquenched with particle disruption (iv). (B) Lipid film hydration/activation protocol described in this report to generate lipid microbubbles from the ground-up and (C) integration of one-pot radiolabeling between lipid suspension formation and microbubble activation. This figure was adapted with permission from Rajora et al.¹⁵. Please click here to view a larger version of this figure.

Protocol

1. Preparations of reagents

1. Prepare ammonium acetate buffer (0.1 M, pH 5.5)
   1. Using an analytical balance, weigh 770.8 mg of ammonium acetate onto a weigh paper. Transfer the weighed amount to a clean 250 mL glass beaker.
   2. Add 90 mL of double distilled water (ddH₂O), measured via a graduated pipette, to the beaker. Add a stir bar and place the beaker on a magnetic stir plate to dissolve the ammonium acetate. Stir at a speed that creates a slight vortex but without solution splashing.
   3. Calibrate a pH meter according to instrument instructions using standards of pH 4 and 7. Once calibrated, insert the pH probe into the ammonium acetate buffer.
   4. Add 104 µL of acetic acid to the solution, stir to dissolve, and measure the pH.
      NOTE: The pH should be close to 5.5 at this point.
   5. Adjust the pH of the buffer by adding 10 N sodium hydroxide (or hydrochloric acid if the buffer becomes too basic) in 5-10 µL increments using a micropipette. Stir, measure the pH, and repeat as necessary. Make a note of the volume of base/acid added.
   6. Add enough volume of ddH₂O to create a total of 100 mL of buffer.
      NOTE: For example, if 45 µL of 10 N sodium hydroxide was used during pH adjustment, 9.851 mL of ddH₂O would be added to the beaker (100 mL [target volume]- 90 mL [step 1.1.2] - 0.104 mL [step 1.1.4] - 0.045 mL [step 1.1.5] = 9.851 mL).
   7. Stir the buffer one final time thoroughly before transferring it to a lidded storage container.
   8. Clean the pH meter as per instrument instructions.
      CAUTION: Concentrated aqueous sodium hydroxide and hydrochloric acid can cause skin reactions and should be handled using gloves.
2. Prepare hydration buffer (PGG)
   1. Aspirate phosphate-buffered saline (PBS) into a syringe and equip the end with a polyethersulfone 0.2 µm pore size syringe filter. Filter the PBS into a clean plastic, lidded centrifuge tube.
      NOTE: 0.2 µm pore syringe filters of alternative membrane materials (for example, polyvinylidene fluoride) can be used so long as the membrane is compatible with PBS and ammonium acetate.
   2. Combine filtered PBS, propylene glycol, and glycerol via micropipette in an 8:1:1 volumetric ratio to make the hydration buffer (also referred to as PGG). When adding propylene glycol and glycerol, aspirate and wipe any residual droplets of propylene glycol or glycerol from the surface of the pipette tip before slowly pipetting the reagent into the PBS. A clear string-like viscous turbidity will be seen in the PBS.
      NOTE: It is recommended to use a p1000 micropipette to first add PBS to a centrifuge tube, followed by propylene glycol and glycerol, as the latter two reagents are viscous. As such, they should be slowly aspirated via the micropipette until fluid movement is no longer seen in the pipette tip and such that no air is uptaken when the pipette tip is removed from the reagent. Micropipette tips with volumetric markings should ideally be used to choose reagent volumes that align with such markings (for example, making 1 mL or 5 mL PGG, and respectively using the 0.1 mL or 0.5 mL marking on the micropipette tip to visualize complete aspiration of propylene glycol and glycerol). When wiping the surface of the micropipette tip, do not wipe at the tip opening, only at the sides.
   3. Pipette up and down with the pipette tip in the solution until the reagents have been homogenously dissolved. Be careful not to introduce any air bubbles into the solution.
   4. To further ensure a complete mixture of the hydration buffer, cap the centrifuge tube and rotate up and down slowly. Do not vortex.
   5. Spin the tube at less than 1000 x g for 20-30 s (minimum temperature 4 °C, maximum RT) to remove unobservable air bubbles.
3. Prepare hydration/radiolabeling buffer (AA-PGG)
   1. Syringe filter 0.1 M, pH 5.5 ammonium acetate buffer (from step 1.1), and PBS into separate tubes as per step 1.2.1.
   2. Combine filtered ammonium acetate buffer, filtered PBS, propylene glycol, and glycerol via a p1000 micropipette into a centrifuge tube in a 5:3:1:1 ammonium acetate buffer: PBS: propylene glycol: glycerol volumetric ratio in the listed order. Follow aspirating, mixing, and centrifugation instructions as per steps 1.2.2-1.2.5 to make AA-PGG.
4. Instant thin layer chromatography (iTLC) eluent
   1. Weigh up to 0.1 g of ethylenediaminetetraacetic acid (EDTA) and transfer to a capped vial. Dissolve in ddH₂O such that a 2% w/v solution of EDTA is made (for example, for 50 mg of EDTA, add 2.5 mL of ddH₂O).
   2. Combine the 2% w/v EDTA solution with the ammonium acetate buffer from step 1.1 in a 9:1 v/v ratio (90% EDTA solution, 10% ammonium acetate buffer). Cap and store the resulting iTLC eluent.

2. Formation of lipid films

NOTE: This procedure outlines the formation of a lipid film with compositions mimicking the commercial microbubble, Definity^®, with porphyrin-lipid substituting the host lipid and constituting 30 mol% of the total lipid. However, the radiolabeling protocol can be applied to diverse lipid formulations (C16, C18, C22 chain lengths, neutral or anionic charge, varying porphyrin-lipid molar compositions). A Supplementary Spreadsheet (Supplementary File 1) is attached that provides calculations, compositions, masses and stock volumes for the described and other formulations. All lipids are commercially available with the exception of the porphyrin-lipid, pyropheophorbide-a-lipid (pyro-lipid), the synthesis of which has been previously described in detail^35,36.

1. Using Supplementary File 1, determine the total mass needed for each lipid based on the number of films required.
2. Weigh an empty 0.5 dram glass vial on an analytical balance.
   NOTE: Dust interferes with successful microbubble formation. Thus, blow pressurized air into the vial to remove any dust/particulates if stored uncapped.
3. Weigh 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) onto weigh paper.
   NOTE: The mass weighed should be obtained from step 2.1 plus an additional 0.5-1 mg to account for any loss during sample handling in later steps.
4. Transfer the DPPC to the weighed glass vial and reweigh it to determine the lipid mass in the vial. This process allows for easier lipid transfer to the glass vial, reduced lipid powder loss/spillage, and more accurate measurement of the lipid mass.
5. Repeat steps 2.2 through 2.4 with the other lipids: 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-5000] (DPPE-mPEG), 1,2-dipalmitoyl-sn-glycero-3-phosphate (DPPA), and C16 pyro-lipid.
   NOTE: If pyro-lipid is not available in a weighable powder form but rather as a film or aliquot of unknown quantity, it can be dissolved in chloroform to form a stock whose concentration can be calculated through UV-Vis absorbance measurements in methanol using the Beer-Lambert law as previously described³⁵.
6. Prepare the following organic solvents and solutions in glass test tubes using micropipettes or glass syringes: 1) chloroform, 2) 9:1 v/v chloroform: methanol, and 3) 65:35:8 chloroform:methanol:ddH₂O. For the last, pipette the components and mix them in the following order: ddH₂O, methanol, then chloroform.
   CAUTION: Methanol and chloroform are health hazards, flammable and volatile. Wear eye protection, gloves, and a lab coat, and use a fume hood.
7. Use the Supplementary File 1 to calculate the volume of organic solvent/solution needed to make lipid stocks and select glass syringes of appropriate volumes.
   NOTE: This volume should yield stock concentrations that correspond with 15-100 µL stock aliquot volumes per film that can be readily measured using 25-100 µL glass microliter syringes.
8. Rinse the glass syringes thrice with chloroform. Pump the plunger back and forth to dry the syringe.
9. Measure and add the organic solvent/solutions via the cleaned glass syringe to the individual lipid vials as per the spreadsheet calculations in step 2.7 to form lipid stocks. Dissolve pyro-lipid in chloroform (unless already dissolved as per the step 2.5 note), DPPC and DPPE-mPEG in 9:1 v/v chloroform: methanol and DPPA in 65:35:8 chloroform:methanol:ddH₂O. If using the same glass syringe for all additions, rinse and dry between each lipid.
   NOTE: If the formulation of choice does not contain DPPA or its C18 chain length variant, then pyro-lipid, host PC lipid, and PEG lipid can all be dissolved in chloroform.
10. Cap the vials and vortex.
11. Add calculated volumes of stock lipid solutions to a new 0.5-dram glass vial (film vial) via a glass microliter syringe. For the first lipid stock, insert the needle tip into the bottom center of the vial and plunge slowly to avoid splashing up the vial walls. For subsequent additions, place the needle tip directly above the liquid level and touch the side of the vial to remove any final drops in a manner that does not expose the needle to the liquid below.
    NOTE: Rinse and dry the glass syringe between lipid additions when contamination occurs. If making multiple films, cap both the film and stock vials between additions to minimize solvent evaporation.
12. Gently swirl the vial manually in an upright position to mix contents. Avoid splashing any solution up the vial walls.
13. Uncap (store the cap) and insert a nitrogen line into the headspace of the vial. Adjust the nitrogen flow to cause a slight visible disturbance at the liquid surface but without any funneling or splashing.
14. Vortex the vial immediately after inserting the nitrogen line. Start at a low speed sufficient to form a funnel with solvent rising no higher than 1 cm from the bottom of the vial. Avoid solvent splashing. As the solvent evaporates, increase the vortex speed slowly and without pause, maintaining the solvent height until all liquid evaporates. The result will be a thin film coated across the lower third of the vial.
15. Place the vial in a vacuum-equipped desiccator and continue to dry the film under vacuum for 8-72 h. Cover the vial (except the opening) or the desiccator with aluminum foil.
    NOTE: The protocol can be paused here. The next steps can be performed after film drying, or the films can be stored under argon, sealed with Parafilm, in a -20 °C freezer for up to 1 month, and longer if kept dry.

3. Lipid film hydration

NOTE: If the microbubbles are used in vitro or in vivo, use sterile micropipette tips, tubes, syringes, and needles for steps 3.3 through 5.4 unless otherwise specified.

1. Remove the film from the vacuum or, if freezer-stored, allow it to warm to RT.
2. Fill a 250 mL beaker with water and heat the water to 70-80 °C.
3. Heat a water bath sonicator to 69 °C.
4. Micropipette 1 mL of AA-PGG (step 1.3) down the edges of the lipid film vial to avoid bubble creation.
   NOTE: When fabricating unchelated control or fluorescent-only microbubbles, use PGG (step 1.2) instead of AA-PGG.
5. Partially cover the vial opening with a cap, leaving enough space to insert a perfluoropropane (PFP) line. Flow PFP into the vial headspace for 20 s above the liquid, such that the liquid is visibly disturbed but does not splash. Do not flow PFP directly into the suspension. Cap the vial.
   NOTE: If the flow is adequate in strength and time, the vial will start to cool to the touch.
6. Submerge the bottom half of the vial into the 70-80 °C water bath for 1 min. Then, sonicate for at least 30 s in the 69 °C bath sonicator or until the lipid film disperses homogeneously into the AA-PGG. Avoid creating bubbles or prematurely activating microbubble formation (premature activation will appear as milky/cloudy areas in the lipid suspension). Wipe the vial surface when needed to better discern if any unsuspended lipids remain.
   NOTE: If the lipid film does not hydrate within 1 min of sonication, re-heat in the 70-80 °C  bath, and re-sonicate.
7. Once the lipid film is homogenously suspended, heat one last time for 1 min and sonicate for an additional 30 s.
8. Wipe the vial and allow it to cool passively to RT (~5-10 min).
9. Refill the vial headspace with PFP as per step 3.5, cap, and seal the cap edges with Parafilm.
   NOTE: The protocol can be paused here and resumed after no later than 8 h.

4. Radiolabeling

NOTE: For unchelated control or fluorescent-only microbubbles, skip to protocol Section 5.

CAUTION: Perform steps 4.4-4.6 of this protocol in a radioactive laboratory unless otherwise specified. ⁶⁴CuCl₂ is a radiological hazard with a risk of multisystem toxicity through skin exposure, inhalation, or ingestion. Whenever possible, handle it in a fume hood indirectly using rubber-tipped forceps. Wear a protective lab coat, a personal ring and badge dosimeter, and double glove when handling. Ensure ⁶⁴CuCl₂ is handled across 2-inch lead shielding. When necessary, transport it in a lead-sheathed container. Shield waste containers and conduct an operational survey for contamination following use.

1. Prepare a 60 °C water bath in a glass beaker or large crystallizing dish containing a magnetic stir bar. Use a temperature-controlled hot/stir plate equipped with a thermal probe inserted into the water, set to stir at a rate that produces a weak but visible funnel.
2. Transfer a sealed vial containing ⁶⁴CuCl₂ in 0.1 N HCl to a dose calibrator via rubber-tipped forceps.
   NOTE: When ordering ⁶⁴CuCl₂, request it be dissolved in 5-20 µL of 0.1 N HCl. A lower volume is critical for preserving microbubble yield.
3. Note the copper-64 activity measured on the dose calibrator and the time. Remove the vial using forceps and place it in a leaded container.
4. Divide the activity noted by the volume reported for the ⁶⁴CuCl₂ to obtain a MBq·mL^-1 value.
5. Uncap the lipid suspension from step 3.9 and secure it in a vial holder.
6. Uncap the ⁶⁴CuCl₂ vial and secure it with forceps.
7. Micropipette a volume of the ⁶⁴CuCl₂ solution corresponding to 40-250 MBq of activity and transfer into the lipid suspension. Ensure the pipette tip is submerged into the suspension. Plunge and then pipette up and down to completely transfer the ⁶⁴CuCl₂.
   NOTE: The amount of ⁶⁴CuCl₂ added will depend on the intended application for the radiolabeled microbubbles and the sensitivity of the dose calibrator. For longitudinal (up to 48 h post-injection) PET and blood sampling in vivo in mice, a minimum of 220 MBq and 50 MBq, respectively, are recommended.
8. Cap both lipid suspension and ⁶⁴CuCl₂ vials.
9. Using flat rubber-tipped forceps, manually rotate the radioactive lipid suspension up and down at least 5 times to gently mix the ⁶⁴CuCl₂ through the suspension. Avoid shaking or dropping the vial, and avoid bubble formation.
10. When right-side-up, gently flick the cap of the vial while keeping the suspension stabilized. This will help any liquid trapped in the cap to gravitate to the bottom of the vial. Carefully partially uncap the vial and insert an 18 G needle-equipped PFP line. Fill the vial headspace with PFP for 20 s as per step 3.5. Cap the vial and seal with Parafilm.
11. Measure the vial activity on a dose calibrator and note the time.
    NOTE: If adequate activity was not transferred to the vial, repeat Steps 4.5-4.11, adding an appropriate additional volume of ⁶⁴CuCl₂.
12. Place the vial in a foam vial holder and push through so the bottom half of the vial will be exposed to heat. Place the holder in the stirring 60 °C water bath and heat for 1 h.
13. While the chelation reaction ensues, prepare iTLC plates. While wearing fresh gloves, cut glass microfiber chromatography paper into 1 cm x 8 cm strips. Heat the strips in an 80 °C glass-drying oven.
    NOTE: This step can be performed in a non-radioactive laboratory.
14. After 1 h, remove the vial in step 4.12 from the heat and wipe the edges with tissue.
15. Rotate the vial up and down manually with rubber-tipped forceps to recondense any condensation on the vial walls into the lipid suspension.
16. With the vial in an upright position, flick the cap while stabilizing the tube. Remove the Parafilm and wipe around the cap to remove any trapped bath water.
17. Carefully uncap the vial and aspirate 1-2 µL of the lipid suspension. Spot the suspension 1 cm from the bottom center of an iTLC strip and re-cap the vial. Allow the spot to dry.
    NOTE: Ideally, a minimum of 2 iTLCs should be spotted per reaction mixture and developed per radiolabeled lipid suspension for certainty.
18. Micropipette 200 µL of the iTLC eluent (prepared in step 1.4) into the bottom of a 10 mL test tube. House the test tube in a lead container. Add the spotted iTLC to the tube and allow the strip to develop until the eluent is approximately 1 cm from the top edge of the strip.
19. Remove the developed iTLC strips using forceps. Hold the strip vertically and cut into thirds over γ-counter and push-cap compatible round-based 5 mL plastic tubes such that each strip third falls directly into an individual third. Insert push caps into the three tubes.
20. Measure the strip-containing tubes and an empty/capped control tube on a γ-counter for copper-64 activity and record the associated counts-per-minute (cpm). Subtract the control tube activity from the other readings to correct for background activity.
    NOTE: The corrected readings for the bottom third of the strip (piece 1) are associated with copper-64 chelated to lipid suspension particles. The middle section (piece 2) contains a streak of free copper-64 and ⁶⁴Cu-pyro-lipid chelates in non-supramolecular form. The top section (piece 3) predominantly contains free copper-64.
21. Calculate radiochemical purity via Equation 1.
         (Equation 1)
    NOTE: if the cpm for piece 1 seems unreasonably low (for example, lower or equivalent to pieces 2 or 3) or if any readings are above the γ-counter non-linear/saturation threshold, spot a lower volume or a diluted aliquot (1-2 µL) of the radiolabeled suspension for iTLC.
22. Ensure that the radiochemical purity obtained from both iTLC strips per lipid suspension is ≥ 94% to continue. If not, continue to heat the lipid suspension at 60 °C and monitor the chelation at 30 min intervals via iTLC.
23. Uncap the radiolabeled lipid suspension vial and micropipette 8.89 µL of 1 N NaOH into the suspension, pipetting up and down to completely transfer the base and neutralize the suspension. Cap the vial, rotate manually with forceps to invert/revert, and then gently tap the vial cap.
24. Fill the headspace with PFP as per step 4.10, cap and seal with Parafilm.

5. Microbubble activation and isolation

1. Activate the lipid suspension via a mechanical vial shaker for 45 s at 4530 rpm to generate a milky microbubble suspension. Allow the vial to passively cool to RT for approximately 10 min. The resulting milky suspension will separate into two layers over time.
   NOTE: The vial contents should look milky after activation. A clearer suspension post-activation is indicative of unsuccessful activation, the contributors of which will be discussed in later sections.
2. Once at RT, gently invert/revert the vial to resuspend the microbubble suspension. Set the vial on a flat surface and wait 2 min prior to decanting to obtain the desired microbubble population as follows:
   1. Equip a 1 mL plastic syringe with an 18 G needle and vent the syringe/needle by aspirating and plunging air in/out. At the 2 min mark, quickly uncap the vial, breaking the Parafilm seal in one motion.
   2. Draw 400-550 µL from the bottom of the vial (target microbubble population), avoiding aspiration of the top foamy layer of larger undesired microbubble populations.
      NOTE: If needed, tilt the vial to one side to collect the end volumes into the syringe to avoid aspirating the foamy/lighter layer.
3. Wipe the edges of the needle carefully to remove any foamy contaminants and transfer the isolated microbubble suspension into a microcentrifuge tube. Cap gently (do not abruptly snap the cap open or closed). This is the final working suspension of radiolabeled microbubbles.
4. Measure the activity of the final microbubble product on the dose calibrator and note the time. Divide this value by the volume of suspension decanted in step 5.2 to obtain an MBq·mL^-1 value to calculate injection volumes depending on the application of interest.
   NOTE: The radiolabeled microbubbles are now ready to use. Section 6 can be executed up to 24 h later. For information on how these radiolabeled microbubbles can be injected and tracked in vivo through multimodal (ultrasound, PET, fluorescence) imaging, refer to Rajora et al.¹⁵.

6. Validating radiolabeling efficiency

1. Resuspend the microbubble suspension through gentle pipetting or vial inversion.
   NOTE: Never vortex a microbubble working product. Vortexing destabilizes microbubble suspensions.
2. Add 10-200 µL of the radiolabeled suspension to a 0.5 mL 30,000 molecular weight cut-off (MWCO) centrifuge filter unit. If using volumes <200 µL, add ddH₂O to the filter unit to constitute a total 200 µL volume. House the filter unit in a compatible microcentrifuge tube and cap.
   NOTE: Conducting a radiolabeling test prior to and separately from any applied in vitro or in vivo use of radiolabeled microbubbles is recommended to ensure successful protocol completion. In this case, a larger volume (e.g., 200 µL) could be used in this step. When the protocol is subsequently used for a treatment session, prepare treatment volumes/injections first, and then conduct section 6 with the remaining radiolabeled microbubble suspension as early as possible.
3. Centrifuge for 10 min at 12,000 x g at RT.
   NOTE: the microcentrifuge should be surrounded by lead shielding.
4. Cut the connection between the microcentrifuge tube and its cap with scissors.
   1. Place the cap in a 20 mL scintillation vial labeled "caps". Transfer the filter unit to a new microcentrifuge tube (tube 2).
   2. Place the first microcentrifuge tube with infranatant into a 20 mL scintillation vial labeled "tube 1". Add 200 µL ddH₂O to the filter unit in tube 2.
5. Centrifuge the filter unit in tube 2 at 12,000 x g at RT.
6. Repeat steps 6.4 and 6.5. Add the tube 2 cap to the "caps" scintillation vial harboring the tube 1 cap. Place tube 2 in a new 20 mL scintillation vial.
7. Cut the cap of the third microcentrifuge tube and place in the "caps" vial as per step 6.4. Transfer the filter unit to a new 20 mL scintillation vial labeled "unit", ensuring that the infranatant remains in tube 3 and is not transferred to the "unit" vial. If drops are seen on the edges of the filter unit, return it to tube 3, cap, and spin down for 10 s. Place tube 3 in a new 20 mL glass scintillation vial.
8. Cap the 5 scintillation vials (tube 1, tube 2, tube 3, caps and unit). Prepare one empty and capped 20 mL scintillation vial as a blank control.
9. Measure the six scintillation vials on a γ-counter for copper-64 activity. Subtract the blank vial activity from that of other vials. Calculate the radiolabeling/chelation efficiency using Equation 2.
       (Equation 2)
   NOTE: If the unit cpm is unreasonably low (ex, lower or equivalent to the tubes) or if any readings are above the γ-counter non-linear/saturation threshold, store the scintillation vials in lead containers for up to 4 days to allow the activity to decay until values are below the threshold and remeasure.

7. Microbubble physicochemical characterization

NOTE: Unless a laboratory has designated equipment for radioactive sample processing, microbubble physicochemical characterization must be conducted using non-radioactive, "cold" copper-chelated samples. This "cold" labeling facilitates the assessment of microbubble yield, which is vital for assessing the dose of microbubbles used for one's intended application. Additionally, it allows for comparison with control unchelated microbubbles to ensure the radiolabeling process does not perturb the properties of microbubbles. This "cold" labeling and associated physicochemical characterization should occur prior to radiolabeled microbubble application and can be used as feedback if modifications to radiolabeling are required (see Discussion).

1. "Cold" copper microbubble labeling
   1. Using the volume of ⁶⁴CuCl₂ solution added to the lipid suspension in step 4.7, porphyrin molar% composition within the lipid films and specific activity found on the ⁶⁴CuCl₂ product sheet, calculate the approximate metal:porphyrin molar ratio achieved during radiolabeling. Example calculations can be found in Supplementary File 1.
   2. Follow sections 1-3 of the current protocol.
   3. Prepare a 0.1 mg·mL^-1 CuCl₂ solution in 0.1 N HCl.
   4. Micropipette the appropriate volume of this CuCl₂ solution into the lipid suspension calculated from step 7.1.1 and cap the vial.
   5. Rotate the vial to mix the CuCl₂ into the lipid suspension, fill the headspace with PFP, seal, and heat as per steps 4.9, 4.10, and 4.12. Rubber forceps are not needed for vial handling.
   6. After 1 h, remove the vial from the heat and wipe the exterior to dry. Allow the vial to cool to RT.
   7. Neutralize the lipid suspension, mix, fill the headspace with PFP, and seal as per steps 4.23 and 4.24.
   8. Activate the microbubble suspension and decant to obtain a working product as per steps 5.1-5.3.
2. Microbubble sizing
   NOTE: Microbubble sizing should be conducted immediately after activation. If assessing the stability of the working suspension, repeat sample preparation and measurements at 30 min intervals. Typically, a 1-2 h window is representative of the timeframe over which the microbubble working suspension would be used/administered post activation. The aim of stability measurements is to ensure microbubble size and yield are maintained across this timeframe so that all treatments administered from the working solution contain similar microbubble populations.
   1. Turn on the Coulter Counter (CC) and set the following parameters using the Edit SOP tool: 30 µm aperture, 0.6-18 µm size range, aperture current 400-600 µA, preamp gain 4-8, 400 bins, flush before and after each run, volumetric analysis, 5 µL sample volume.
   2. Filter CC electrolyte through a 0.2 µm media vacuum filtration unit. Fill the electrolyte container and a separate container for sample preparation.
   3. Background measurement: Fill a 10 mL disposable cuvette with 10 mL filtered electrolyte and run a baseline measurement. Ensure the counts are below 400. If not, flush the instrument.
   4. Sample measurement
      1. Add 10 mL of filtered electrolyte to a new cuvette. Resuspend microbubble suspension by manually inverting/reverting. Micropipette 5 µL from the bottom center of the vial. Wipe the edges of the pipette tip (except the opening) and plunge the sample directly into the prepared electrolyte.
      2. Pipette up and down to completely transfer the suspension. Use the pipette tip to gently swirl the electrolyte until the "wisps" of microbubble suspension are dispersed.
   5. Measure the sample on the CC (two runs per analyte).
      NOTE: A 5 µL microbubble sample volume is typically appropriate for samples containing 1-5 x 10⁹ microbubble·mL^-1 concentrations. This sample volume may need to be adjusted depending on the specific CC instrument setup and if the sample microbubble yields fall outside the above range.
3. Confocal imaging
   NOTE: Conduct confocal imaging immediately after microbubble sizing and within the timeframe of retained microbubble stability as per the note in step 7.2.
   1. Resuspend the microbubble suspension and transfer 1-5 µL to the center of a glass microscope slide. Place a cover slip carefully over the microbubble suspension droplet, avoiding any entrapment of air bubbles. The suspension will spread under the cover slip.
   2. Image the microbubbles at 60x magnification with an oil immersion objective. Obtain images in brightfield and under 633 nm excitation/640-765 nm emission. Overlay the brightfield and fluorescence images.
      NOTE: Fluorescence signal should overlap across the shell of all visible particles when the probe is homogenously incorporated within the microbubble shell.
4. Spectrofluorometry
   NOTE: Spectrofluorometry measurements can be conducted within 24 h of microbubble activation.
   1. Prepare 1% Triton X-100 as previously described³⁵.
   2. Turn on the spectrofluorometer 15-30 min prior to the first measurement.
   3. Measure the fluorescence spectra of 1% Triton X-100 using a 410 nm excitation and 600-800 nm emission range in a quartz cuvette. Select the option to normalize the signal by the reference detector signal (often referred to as S1/R1).
      NOTE: Triton X-100 easily bubbles when pipetted. As such, when transferring to a cuvette, only plunge to the first micropipette stop.
   4. Rinse the cuvette with methanol and dry with pressurized air in between samples.
   5. Transfer 6 mL of 1% Triton X-100 to a 15 mL centrifuge tube. Resuspend the microbubble suspension and aspirate 1 µL via micropipette. Wipe the edges of the pipette tip except the opening and transfer the sample to the prepared 1% Triton X-100, pipetting up and down to complete the transfer. Vortex the solution and transfer it to a quartz cuvette.
      NOTE: Adjust the ratio of sample: 1% Triton X-100 according to instrument sensitivity and non-linear saturation threshold.
   6. Measure this sample using the parameters in step 7.4.3. This measurement corresponds to "disrupted" particles.
   7. Repeat steps 7.4.3-7.4.6 using PBS. This measurement corresponds to "intact" particles.
   8. Baseline-correct the disrupted and intact sample spectra using 1% Triton X-100 and PBS measurements, respectively.
   9. Calculate quenching efficiency (QE) via Equation 3 using integrated baseline-corrected fluorescence signal of the intact sample in PBS (F_PBS) and in 1% Triton X-100 (F_Tx):
          (Equation 3)
5. UV-Vis spectroscopy
   NOTE: Spectroscopy measurements can be conducted up to 72 h after microbubble activation.
   1. Sonicate an aliquot of the microbubble suspension in a microcentrifuge tube using a bath sonicator at RT until the suspension is transparent. This reduces scattering effects during spectroscopy.
   2. Turn on the spectrophotometer 10 min prior to the first measurement. Select a scanning interval of 0.25 nm and a 200-800 nm acquisition range. Enable baseline subtraction.
   3. Use a 1 cm path-length quartz cuvette for measurements. Rinse the cuvette with methanol between measurements and dry with pressurized air.
   4. Obtain a baseline measurement of methanol.
   5. Vortex the sonicated, transparent microbubble suspension and transfer 10-50 µL into a microcentrifuge tube containing 200-1000 µL of methanol. Ensure methanol volume is measured and added to the tube via a clean glass microliter syringe. Vortex the solution to obtain a "disrupted" sample.
      NOTE: Sample dilution will depend on porphyrin loading efficiency and molar% composition. A 20x dilution is appropriate for a 30 mol% pyro-lipid microbubble composition.
   6. Collect UV-Vis spectrum.
   7. Repeat steps 7.5.4 through 7.5.6 using PBS instead of methanol.
      NOTE: A micropipette can be used to measure PBS volumes instead of a glass microliter syringe.

8. Modifications to protocol

1. Alternative chelator
   1. Prepare lipid films as per section 2, substituting the pyro-lipid for an alternative lipid-conjugated copper chelator (for example, 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-diethylenetriaminepentaacetic acid (ammonium salt), referred henceforth as DTPA-lipid). Use the Supplementary File 1 to calculate the mass and stock volumes required.
      NOTE: Testing various molar compositions of an alternative chelator is likely needed to determine the upper limit beyond which stable microbubbles with high yields cannot be generated.
   2. Follow sections 3 through 6 to generate and characterize radiolabeled microbubbles.
   3. Follow steps 7.1 through 7.3 to characterize "cold" copper-chelated microbubbles. Only brightfield confocal microscopy image acquisition is required to assess particle morphology. If the alternative chelator is fluorescent, conduct confocal microscopy with associated excitation and emission wavelengths in addition to steps 7.4 and 7.5.
2. Alternative fluorophore
   1. Prepare lipid films as per section 2, substituting the pyro-lipid for an alternative lipid-conjugated or intercalating fluorophore (e.g., DiI). Use Supplementary File 1 to calculate the mass and stock volumes required.
      NOTE: Testing various molar compositions of an alternative fluorophore is likely needed to determine the upper limit beyond which stable microbubbles with high yields cannot be generated.
   2. Follow section 3 using PGG instead of AA-PGG.
   3. Complete steps 5.1 through 5.3 and steps 7.2 through 7.5.
3. "Spiking" approach: Labeling pre-formed microbubble lipid suspensions
   1. Generate a porphyrin-lipid film as per section 2, using only pyro-lipid with no other lipid constituents. Refer to the Supplementary File 1 for pyro-lipid quantities.
   2. Hydrate the pyro-lipid film as per section 3 using 100-200 µL of AA-PGG (or PGG if radiolabeling is not required) instead of 1 mL of hydration buffer.
   3. Transfer the entire pyro-lipid suspension to a pre-made lipid microbubble suspension.
   4. Fill the headspace with PFP, heat, and sonicate the suspension and seal under PFP as per steps 3.4 through 3.9. Monitor for dispersion of the pyro-lipid suspension into the pre-made lipid microbubble suspension and conduct heat/sonication cycles until completely dispersed.
      NOTE: If using a septum-sealed commercial microbubble vial, the pyro-lipid suspension can be introduced to the vial through a syringe/needle without refilling the vial headspace with PFP.
   5. Conduct radiolabeling (see step 8.3.5.1 below), activation, isolation (see NOTE below), and associated characterization as per sections 4 through 7.
      1. An alternative approach is to first radiolabel the hydrated pyro-lipid suspension, monitor for >94% radiochemical purity, neutralize with 1 N NaOH (modify the volume according to AA-PGG volume used to hydrate the pyro-lipid film), and then introduce the radiolabeled pyro-lipid suspension into the pre-made microbubble lipid suspension as per step 8.3.3.
         NOTE: This modified approach should only be used to generate fluorescent or multimodal radiolabeled microbubbles if followed by an isolation process that removes submicron multilamellar vesicles created during lipid hydration but not incorporated into microbubbles. See Discussion for further details.

Results

The key quantifiable results when fabricating radiolabeled microbubbles are radiochemical purity and radiolabeling efficiency. This protocol uses iTLC and a validated centrifugal procedure, respectively, to characterize each. Figure 2A shows that average radiochemical purities and efficiencies of ≥95% were achieved across commercial microbubble mimicking formulations in which the host lipid was substituted for pyro-lipid at compositions of 1 mol%, 10 mol%, or 30 mol% of the total lipid. The ...

Discussion

The current lipid microbubble radiolabeling protocol achieves >95% radiochemical purity, >95% chelation efficiency, and retention of microbubble physicochemical properties without necessitating any post-labeling purification. These accomplishments represent advancements previously unattained for existing labeling protocols. Lack of purification steps allows quicker use of radioisotopes (in this case, copper-64), and thus, reduction of inefficient activity loss from radioactive decay. The resulting retention of mi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
64CuCl2	Washington University School of Medicine, Mallinckrodt Institute of Radiology	N/A	Order in small volume (<10 µL) dissolved in 0.1 N HCl
Acetic acid	Any company		≥ 95% purity
Aluminum foil	Any company
Ammonium acetate	Any company		Purity: ≥ 98%
Balance - analytical	Any company		Able to measure down to 0.1 mg
Bath sonicator	Any company		Can be heated to 69 oC
CC aperture - 30 micron	Beckman Coulter	A36391	Particle diameter range: 0.6-18 um
CC electrolyte	Beckman Coulter	8546719	Isoton II diluent
CC Software	Beckman Coulter	Multisizer 4e
Centrifuge filter units (0.5 mL 30,000 MWCO) with compatible microcentrifuge tubes	MilliporeSigma	UFC503096	Amicon Ultra - 0.5 mL
Centrifuge tubes - 15 mL with caps	Any company
Chloroform	Any company		Purity: ≥ 99.8%
Coulter counter	Beckman Coulter	B43905	Multisizer 4e Coulter Counter
Cover slips	VWR	48393081	VWR micro cover glass
CuCl2	Any company		Ensure not oxidized
CuCl2
Cuvette- quarts, 1 cm path length	Any company
Cuvettes - 10 mL plastic for CC measurements	Beckman Coulter	A35471	Coulter Counter Accuvette ST
ddH2O	Any company		Can be obtained through an ultrapure water purification system
DiI (1,1'-Dioctadecyl-3,3,3',3'-Tetramethylindocarbocyanine Perchlorate)	Any company		Powder form
Dose calibrator	Any company		Able to read copper-64
DPPA (1,2-dipalmitoyl-sn-glycero-3-phosphate (sodium salt))	Avanti Polar Lipids	830855P	Powder form
DPPC (1,2-dipalmitoyl-sn-glycero-3-phosphocholine)	Avanti Polar Lipids	850355P	Powder form
DPPE-MPEG (1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-5000] (ammonium salt))	Avanti Polar Lipids	880200P	Powder form
DTPA-lipid (1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-diethylenetriaminepentaacetic acid (ammonium salt))	Avanti Polar Lipids	790106P	Powder form
EDTA (Ethylenediaminetetraacetic acid)	Any company
Gamma counter	Any company		Able to read copper-64
Gamma counting tube push caps	Globe Scientific	22-171-665	Flanged plug caps for 12 mm tubes
Gamma counting tubes	Sarstedt	55.1579	5 mL, 75 x 12 mm, PS
Glass beaker - 250 mL	Any company		Able to withstand temperatures up to 100 oC
Glass drying oven	Any company		Can be heated to 80 oC
Glass microliter syringes - 25, 50, 100, 1000 µL	Any company		Compatible with organic solvents
Glass scintillation vials - 20 mL	VWR	66022-081	VWR® Scintillation Vials, Borosilicate Glass, with Screw Caps, With pulp foil liner
Glass vials - 0.5 dram	VWR	66011-020	VWR Vial 1/2 dram, with black phenolic screw cap and polyvinyl-faced pulp liner
Glycerol	Sigma Aldrich	G7757-1L	Purity:  ≥ 99.0%
Graduated pipette/gun	Any company
Hot/stir plate			Equipped with temperature prob for automatic tempearture control
Hydrochloric acid - 0.1 N	Any company
iTLC plates	Agilent	A120B12	iTLC-SA chromatography paper
Laboratory tissues	Any company
Media vaccuum filtration unit	Any company		0.22 micron pore size, PES membrane, 500 mL funnel capacity
Methanol	Any company		Purity:  ≥ 99.8%, HPLC grade, meets ACS specifications
Microcentrifuge tubes non sterile - 1.5 mL	Any company
Microcentrifuge tubes sterile - 1.5 mL	Any company
Micropipetes - p1000, p200, p20, p10	Any company		Ensure are calibrated
Microscope slides	Fisher Scientific	12-550-15	Superfrost Plus Microscope Slides Precleaned
Needles - 18 G			Sterile
Parafilm	Any company
PBS	Sigma Aldrich	D8537-500ML	DPBS, modified, without calcium chloride and magnesium chloride, liquid, sterile-filtered, suitable for cell culture
PFP	FluoroMed	APF-N40HP	Purity:  ≥ 99.8%
PFP line	Any company		1/4 inch diameter plastic hose cut about 50 cm in length
PFP regulator	Swagelok	SS-1RF4 and SS-4HC-1-4
pH meter	Any company
pH standards 4 and 7	Any company
Pipette tips for p1000, p200, p10 - non sterile	Any company
Pipette tips for p1000, p200, p10 - sterile	Any company
Plastic syringe - 1 mL	Any company		Sterile
Propylene glycol	BioShop	PRO888.500	Purity:  ≥ 99.5%
Pyro-lipid	N/A		Made in-house
Rubber tipped forceps	Any company		Mix of fine-tipped and flat/square edges recommended
Scissors	Any company
Sodium hydroxide - 1 N	Any company
Sodium hydroxide - 10 N	Any company
Spectrofluorometer	Any company		Capable of 410 nm excitation and 600-850 nm emission
Spectrofluorometry software	Horiba	FluorEssence
Spectrometer	Any company
Syringe - 1 mL	Any company		Disposible, plastic, sterile
Syringe filters - 0.2 micron pore size	Any company		Membrane material: PES or other compatible with ammonium acetate/acetic acid and PBS
Test tube - 10 mL
Triton X-100	Any company
Vacuum desicator/vacuum	Any company
Vialmix	Lantheus Medical Imaging	515030-0508	Referred to in protocol as a mechanical vial shaker
Weigh paper	Any company		To avoid losing product, cutting weigh paper into 3x3 cm squares is recommended