Liposomal local anesthetics increase the duration of drug presence at target sites,1,2 thus prolonging analgesic effect.3 Reduced plasma redistribution causes lower peak plasma drug concentrations (Cmax),4 which, in turn, allows the safe administration of larger drug doses, further prolonging the analgesic effect.4 Currently available liposomal local anesthetic preparations are limited by complicated and expensive manufacture and liposomal loading, and by the leakage of drug from the liposomes, leading to short shelf-life despite refrigeration (<1–2 months).2,5,6
Herein, we describe the preclinical development of a novel formulation of proliposomal ropivacaine, which is an aqueous-free, oil-based ropivacaine formulation that leads to liposomal formation only on contact with aqueous subcutaneous tissue. Specifically, we report the following: (1) the manufacture of the novel formulation, (2) the conversion of the study drug from a particle-free homogenous oil to an emulsion containing multilamellar liposomal vesicles after exposure to saline or plasma as models of extracellular fluid, (3) the demonstration of extended drug shelf-stability for at least 2 years, (4) local toxicity data in pigs, and (5) pharmacodynamic and pharmacokinetic data in piglets. In an accompanying report, we describe pharmacodynamic and pharmacokinetic data in healthy human volunteers after subcutaneous proliposomal ropivacaine administration.7
All in vivo experiments were performed at a limited-access pig facility by MDBiosciences, Ltd., in the Institute of Animal Research, Lahav, Israel, in accordance with the guidelines and approval of the Institutional Animal Care and use Committee. Animals were provided with a commercial pig diet and had free access to drinking water ad libitum. All animals were under veterinary supervision and were assessed daily by an investigator for signs of distress and to ascertain that they were eating and drinking normally.
Study Drug: Preparation
Proliposomal 4% ropivacaine was prepared by Nextar Chempharma Ltd., Israel, from the following components: ropivacaine HCl H2O (Haorui Pharma-Chem Inc., China), lecithin phospholipon 90G (PL90G; Phospholipid GmbH, Germany), castor oil (Spectrum Chemical MFG Corp., New Brunswick, NJ), cysteine HCl (Spectrum Chemical MFG Corp.), absolute ethanol (EtOH; Spectrum Chemical MFG Corp.). All components met GRAS criteria (“generally regarded as safe”) of the Food and Drug Administration. The composition of proliposomal 4% ropivacaine was ropivacaine HCl monohydrate base 4.78 % (w/w), PL90G 53.91 % (w/w), castor oil 35.21 % (w/w), cysteine HCl 0.10 % (w/w), and ethanol 6.0 % (w/w). Ingredients were weighed and dissolved in ethanol and blended by sonication in a pharmaceutical reactor at 50°C. Sonication was continued for 4 hours and additional 1-hour periods, if needed, until a clear solution was obtained. The solution was dried in a rotary evaporator (rotor 60 rpm, vacuum 40–200 mbar), and final alcohol content was corrected to 6%.
Study Drug: Development of Multilamellar Liposomal Vesicles
We assessed the effect of 2 models of aqueous subcutaneous tissue (saline and plasma) on the development of nanoparticles in the proliposomal ropivacaine preparation. The formulation was assessed for nanoparticles using a particle-size distribution analyzer, light microscopy, and cryo-transmission electron microscopy (cryo-TEM).
In Vitro Simulation with Saline
Proliposomal ropivacaine was divided into 4 scintillation vials; 0.9% saline solution was slowly injected into the formulation in quantities of 10%, 20%, 40%, and 50% (w/w). Vials were lightly agitated at 37°C using a water bath shaker. A turbidity test was performed against a blank water sample using an Agilent 8453 UV-visible Spectrophotometer (Agilent Technologies, Santa Clara, CA). The sample was scanned in the visible range of 400 to 700 nm. Sample was diluted with water before the analysis to achieve an absorption value below 1.0. Sedimentation was determined by centrifugation at 15,000 rpm for 10 minutes. Particle size distribution was analyzed using the Coulter LS230 Particle Size Analyzer (Beckman, Brea, CA). Further estimation of the lipid dispersion droplet sizes within the micrometer scale range used light microscopy (Nikon Eclipse E600, Japan) with Coulter Instrument Standard 15-μm Garnet particles as a reference standard. Samples were tested for macroscopic and microscopic appearance at several time points during the incubation period.
In Vitro Simulation with Plasma
Proliposomal ropivacaine was diluted with minipig plasma at dilutions of 1:1 and 1:2 (drug:plasma). Samples were incubated at 37°C for 1 and 5 hours and stirred gently at 50 rpm. For cryo-TEM analysis, a drop of sample was placed on a carbon-coated polymer film supported on a 300 mesh Cu grid (Ted Pella Ltd., Redding, CA), and excess liquid was blotted. Specimens were vitrified by fast quench in liquid ethane to −170°C (Vitrobot, FEI, Hillsboro, OR). Vitrified specimens were transferred into liquid nitrogen for storage. Cryo-TEM used a FEI Tecnai 12 G2 electron microscope (120 kV) with a Gatan cryo-holder maintained at −180°C; images were recorded on a slow scan-cooled charge-coupled device camera (Gatan Inc., Pleasanton, CA). Images were recorded with Digital Micrograph (software, Gatan Inc.) at low-dose conditions to minimize the electron beam radiation damage.
Study Drug: Shelf-Stability
The proliposomal ropivacaine formulation was tested for stability of the ropivacaine assay (see below) over 24 months at normal conditions (25°C, 60% humidity) and over 18 months at accelerated conditions (40°C, 75% humidity). The formulation was also tested for bacterial growth using the standard limulus amebocyte lysate test, with results expressed in endotoxin units/mL (EU/mL).
Local Toxicity: Surgical Wound Model
Ten young piglets (9–11 kg) were randomly assigned to proliposomal 4% ropivacaine (n = 4), plain 0.5% ropivacaine (n = 4), and sham controls (n = 2). Under isoflurane anesthesia and sterile surgical conditions, a single 6- to 7-cm-long incision was made through skin and fascia in the left flank. The wound was closed by continuous suture, and 2.5 mL of the study drugs were administered by subcutaneous injection into the fascial pockets on either side of the wound (total 5 mL). Wound care included chloramphenicol ointment and IM antibiotic administration (50 µg marbocyl and 100 µg amoxicillin). Surgical wounds were photographed and inspected clinically, every day for 2 weeks and assessed histologically from postmortem specimens at 2 weeks.
Two large adult pigs (61 and 67 kg) were also assessed in a multiple-incision wound-healing model. There were 2 incisions for each of the 4 treatment interventions: proliposomal 4% ropivacaine, plain 0.5% ropivacaine, proliposomal vehicle, and sham uninjected control. For each treatment intervention, 1 of the 2 incisions received 3 mL subcutaneous injections to the fascial pocket below the closed surgical wound, as described earlier, and the other incision received direct intraoperative instillation to the open wound. In a ninth site, 3 mL proliposomal 4% ropivacaine was administered subcutaneously to intact skin with no incision. Surgical wounds and injection sites were inspected and assessed clinically and histologically as described earlier.
Seventeen young piglets (11–12 kg) not involved in the earlier studies were randomly allocated to proliposomal 4% ropivacaine (n = 6), plain 0.5% ropivacaine (Astra-Zeneca, Södertälje, Sweden; n = 5), and sham (n = 6) study groups. During a 5-day prestudy acclimation period and throughout the study, 2 investigators hand-fed the animals twice daily and spent time playing with them to habituate them to human handling. Environmental conditions were continuously monitored and carefully controlled: temperature 20°C to 24°C, relative humidity 30% to 70%; 12:12 hour light:dark cycle; 15–30 air changes per hour.
The pharmacodynamic study assessed tactile threshold for mechanical static stimuli using calibrated von Frey filaments (Stoelting Co, Wood Dale, IL) ranging from 1 to 60 g/mm2. Filaments were applied over the surgical wound for approximately 1 second. The different von Frey filaments were applied 3 to 5 times to the tested flank in an ascending order of stiffness, using the “up–down method.”8 Tactile threshold was defined as the smallest force (gram per square millimeter) necessary to cause flank withdrawal (either moving away or twisting away). Study protocol required that baseline tactile threshold be elicited ≥26 g/mm2. Hyperalgesia was defined as a flank withdrawal at a force ≤10 g/mm2. Scores were censored if tactile threshold exceeded 60 g/mm2. In all groups, von Frey filament sensory assessment was performed at baseline before surgery and at 1.5, 3, 6, 8, and 12 hours postoperatively. Animals with persistent analgesic effect at 12 hours were also tested at 18, 24, 30, and 33 hours. Sensory testing for all animals was performed, whereas the pigs were being hand fed by the technician to whom they were accustomed. Sensory testing was also performed on the contralateral side throughout the study period to confirm stable sensory data over time. The area under the tactile threshold versus time curve (AUC) was determined. The AUC between any 2 data points was calculated by using linear interpolation (or the trapezoid rule) as determined by the following formula: AUC = (E1 + E2)/2 × (T2−T1), where E1 and E2 are effect measures at successive data time points, and T1 and T2 are the hours from initial drug administration. The total AUC was determined by the sum of all the component AUCs.
Pharmacokinetic Study Protocol
Eight young piglets, not involved in the earlier studies, were randomly allocated to proliposomal 4% ropivacaine (n = 4; mean ± SD weight, 11.7 ± 0.52 kg) and plain 0.5% ropivacaine (n = 4; mean ± SD weight, 12.2 ± 0.55 kg) injection groups. Investigators performing the study were not blinded to group allocation, but the technician performing the ropivacaine assay was blinded. Drug was injected subcutaneously as two 2.5-mL injections into the surgical incision as described earlier. Total drug dose was 200 mg proliposomal ropivacaine or 25 mg plain ropivacaine. After drug administration, blood was sampled from an ear vein in unanesthetized animals to determine the plasma ropivacaine concentration at baseline and at 0.5, 1, 1.5, 2, 3, 6, 9, 12, 18, 24, 30, 36, and 48 hours in the proliposomal ropivacaine group and at 0.5, 1, 1.5, 2, 3, 4, 5, 6, 9, 12, and 24 hours in the plain ropivacaine group. A paired sample of wound exudate and plasma was also obtained at 96 hours from animals in both groups before euthanasia.
Plasma Sampling and Handling
Venous blood was aspirated from a 16-gauge IV catheter after the aspiration of 10 mL of dead space and collected into cooled EDTA vials. The vials were gently vortexed to ensure that all blood came into contact with the test tube wall. Samples were immediately placed on ice and centrifuged within 15 minutes of sampling in a cooled centrifuge (4°C) at 2000 rpm. Plasma was separated, and samples of 2 mL plasma were labeled and stored at −70°C.
Ropivacaine assay was performed using high-performance liquid chromatography using Waters 2690 separation module and Waters 996 PDA detector (Waters, Milford, MA). The best chromatographic conditions for which retention time, peak shape, and signal-to-noise ratio were optimal were obtained using an isocratic mode with a Synergi column (150 × 2 mm, 4 µm, Polar-RP, 80 Å, Phenomenex, (Torrance, CA). The mobile phase was composed of acetonitrile:2.5 mM ammonium acetate water solution:formic acid in a ratio of 40:60:0.2, v:v:v. The flow rate and the column were maintained at 0.35 mL/min and at 40°C, respectively. high-performance liquid chromatography mass spectrometry analysis was performed with an Agilent 1100 series system (including a quaternary pump, a degasser, and a thermostated autosampler) coupled with a triple quadrupole Qtrap 3200 Turbolon Spray detector (MDS Sciex, Canada) in a positive ionization mode. Detection and quantitation were carried out by multiple reaction monitoring with transitions from m/z 275.4 to 126.1 and 288.9 to 140.2 for ropivacaine and internal standard (bupivacaine), respectively. One hundred microliters of plasma (spiked with ropivacaine) were treated with 200 µL of 4 ng/mL internal standard (bupivacaine) solution in acetonitrile. After agitation and ultracentrifugation, 100 µL of the supernatant were diluted with 100 µL of 0.2% formic acid solution in water, and then 20 µL of this mixed solution were injected in the chromatographic system. Under these conditions, the retention times of ropivacaine and its internal standard (bupivacaine) were 2.1 and 2.6 minutes, respectively. The total run time was 4 minutes. The limit of quantitation was 50 pg/mL for ropivacaine. The signal-to-noise ratio was 16 at the limit of quantitation. Linearity was obtained (linear regression with weighing 1/x2) up to 20,000 pg/mL (r = 0.998). The linearity of the calibration curve ranged between r2 = 0.993 and 0.999 over 4 consecutive analysis dates for the range of 0.2 to 209 ng/mL ropivacaine, and the accuracy of quality control samples ranged from 94.4% to 114.6%.
Data were presented as mean ± SE. The primary end point was the AUC of von Frey tactile threshold over time, which was compared between proliposomal 4% ropivacaine and plain 0.5% ropivacaine using Student t test for all pairwise comparisons with Bonferroni adjusted P values. Statistical significance was assumed at P ≤ 0.01. Statistical analysis was performed using SPSS version 20.0 (IBM Corp., Armonk, NY). No a priori sample size calculation was performed for this proof of concept experimental study outcome. In post hoc power analysis, a conservative approach using the largest within-group SD (271.0, proliposomal ropivacaine) among the 3 groups resulted in tactile anesthesia AUC effect size of 1.295, with corresponding power of 98.1% (α = 0.05). Sensitivity analysis used the smallest within-group SD results in β = 1.00 (G*Power software, v 126.96.36.199, Universitat Dusseldorf, Germany).
The concentration-time data obtained in the study were analyzed using both noncompartmental and compartmental approaches as appropriate. The following pharmacokinetic parameters were calculated: peak plasma ropivacaine concentration (Cmax), the time to achieve the maximum plasma concentration (Tmax), the terminal half-life, and the area under the ropivacaine time-concentration curve extrapolated to infinity (AUC). Pharmacokinetic data were analyzed using WinNonlin software (WinNonlin Version 4.5, Pharsight Corp., Mountain View, CA).
Study Drug: Development of Multilamellar Liposomal Vesicles
Before its exposure to aqueous solutions, the proliposomal ropivacaine formulation was particle free, and no liposomes or any lamellar structures were observed. In vitro exposure to either saline or plasma caused the formation of an emulsion containing multilamellar vesicles.
There was a dose–response effect of aqueous media on liposome formation. Increasing saline content (10%, 20%, 40%, and 50% saline) led to progressively higher macroscopic turbidity and more uniform lipid dispersion (Fig. 1, A–D), with gradual reduction in droplet size leading to multilamellar vesicle formation. There was also a time-dependent effect, with increasing turbidity over the first 7 hours of saline exposure. There was no diminution of turbidity over the next 65 hours.
Particle size distribution analysis (Fig. 1E) demonstrated that mean (SD) particle diameter 1.39 (0.58) μm corresponds more to multilamellar particles (as expected of the multilayer structures) than to micelle particles or oil-in-water emulsion droplets, which are significantly smaller in diameter in the nanometer range.
Multilamellar liposomal vesicles were also produced using minipig plasma as the model for extracellular fluid. These were imaged using cryo-TEM (Fig. 1, F–H).
Study Drug: Shelf-Stability
Ropivacaine concentration was constant throughout the entire period of the assessment (Fig. 2). There was no bacterial growth at any point over 6 months of culture study, and the concentration of bacterial endotoxin at each time point was <10 EU/mL (accepted standard is <100 EU/mL).
Local Toxicity: Surgical Wound Model
Representative histologic and macroscopic appearance of the surgical wounds are shown (Supplemental Digital Content, Supplementary Figs. 1 and 2, http://links.lww.com/AA/B370). All incisions were acceptably healed in both the single-incision piglet models and the multiple-incision large adult pig models. No significant clinical veterinary findings suggested impaired wound healing. In histologic evaluation, the skin gap produced by the surgical incision was bridged by fibrous tissue and reepithelialized in all samples; significant neutrophilic infiltration was not seen at any of the inflamed sites. There was no effect of treatment group on clinical or histologic assessments of wound healing.
One animal in the plain ropivacaine group was excluded because of a surgical wound infection that made sensory assessment at that site impossible. All other animals completed the study as planned.
The von Frey tactile threshold over time is represented in Figure 3. Subcutaneous administration of plain ropivacaine and proliposomal ropivacaine into the closed surgical incision provided effective sensory anesthesia to von Frey stimuli for 6 and 30 hours, respectively. There was an approximately 7-fold increase in the AUC of the individual tactile anesthesia effect–time curve for proliposomal ropivacaine compared with plain ropivacaine (mean difference, 1010; 95% confidence interval [CI], 625–1396 g·h/mm2; P < 0.0001).
The raw data of plasma ropivacaine concentration over time for animals in the different treatment groups (Fig. 4) were assessed by noncompartmental analysis of the data performed using individual plasma concentration-time profiles and doses normalized to subjects’ body weights (Table 1). There was an approximately 5-fold increase in the terminal half-life (±SD) for the proliposomal ropivacaine (16.07 ± 5.38 hours) compared with plain ropivacaine (3.46 ± 0.88 hours); P = 0.0036. There was an approximately 13-fold increase in the Tmax for the proliposomal ropivacaine (6.50 ± 6.35 hours) compared with plain ropivacaine (0.5 ± 0.0 hour).
There was an approximately 8-fold difference in the mean AUC for plain ropivacaine (6.36 ± 2.07 h·mg/L) and proliposomal ropivacaine (47.72 ± 7.16 h·mg/L); P < 0.0001. However, there was also an approximately 8-fold difference in drug dose, so that the dose-normalized mean AUC for plain ropivacaine and proliposomal ropivacaine in our study were not significantly different (3.08 ± 1.00 and 2.77 ± 0.34 h·mg·L-1/mg·kg-1).
The plasma and local wound ropivacaine concentrations 96 hours (4 days) after drug administration are reported for both plain and proliposomal ropivacaine (Table 2). Although proliposomal ropivacaine was associated with an approximately 15-fold higher ropivacaine concentration in plasma at 96 hours (mean difference, 29 ng/mL; 95% CI, 14–44; P = 0.002), this was well below the toxic thresholds.9 By contrast, there was an approximately 250-fold higher concentration in the surgical wound (mean difference, 3783 ng/mL; 95% CI, 1708–5858; P = 0.001), and an approximately 25-fold higher wound:plasma ropivacaine concentration ratio (mean difference, 126; 95% CI, 38–213; P = 0.011).
In this study, we demonstrate the manufacture and preclinical application of proliposomal ropivacaine oil. The shelf-stability of proliposomal ropivacaine was at least 2 years at room temperature. We demonstrate that exposure of proliposomal ropivacaine to aqueous solutions leads to the formation of multilamellar liposomal vesicles in vitro. These vesicles were in the 500 to 2000 nm range, with a SD of vesicle size of ±580 nm, which is similar to the ±544 nm in the previous study using liposomal bupivacaine.10 The pharmacokinetic–pharmacodynamic profile of subcutaneous proliposomal ropivacaine was typical for a liposomal depot preparation4 with a prolonged drug effect, a prolonged Tmax and with a Cmax below the putative toxic threshold (600 ng/mL10) despite a total dose 8 times the maximal allowable total dose for plain ropivacaine.
The dose-normalized mean AUC for plain and proliposomal ropivacaine by subcutaneous administration in our study did not differ, and both were similar to the AUC observed after intraperitoneal administration of plain ropivacaine in pigs.11 However, there was a markedly prolonged tail of plasma redistribution, reflecting the delayed redistribution of ropivacaine from the surgical wound. As no animals received IV ropivacaine in this study, the calculation of absolute ropivacaine bioavailability after subcutaneous administration was attempted based on the existing literature. Although there are some published reports of IV pharmacokinetics of ropivacaine in pigs, these reports studied a very short exposure period (30–60 minutes) and reported much higher concentrations than were observed in our study.12,13 These data could not be used to reliably estimate the AUC after IV administration, and, therefore, the absolute ropivacaine bioavailability in our study could not be calculated. However, another study in rats reported subcutaneous bioavailability of 87% to 130%.14 Taken together with our data in volunteers,4 this information supports the assumption that the bioavailability of subcutaneously administered plain ropivacaine in pigs is close to 100%. The mean concentration-time profile for subcutaneous plain ropivacaine can be fitted with a 2-compartment model with first-order absorption (Table 3 and Fig. 5). However, because of the paucity of data in the absorption phase, the absorption rate constant (K01) could not be estimated with sufficient precision.
Important technical limitations of all liposomal drug therapies involve drug preparation and controlled release. Drug preparation typically includes the preparation of liposomes and active drug loading.10 From our study, it would appear that drug-containing liposomes are generated spontaneously after the oily vehicle is exposed to aqueous tissue, thus avoiding the need for complex and costly liposome manufacture and active drug loading. There are several key stages of drug release from liposomes that should be met for safe and effective controlled release.15,16
1. Before Injection
During storage, there should be no unwanted drug release (leakage) from liposomes, particularly if using liposomal drug doses above the toxic threshold of plain drug.15 However, previously reported liposomal preparations of local anesthetics have high rates of liposomal leakage between 5% and 49%.6 This leakage is both time and temperature dependent6; the only current commercially available liposomal bupivacaine formulation is limited to 30 days at room temperature (20°C–25°C).a With proliposomal ropivacaine, liposomes are only formed on contact with aqueous matter after injection, before which all drug is in the free state.
2. After Injection but Before Arrival at the Site of Action
In situations where liposomes have been used as vehicles to allow systemically administered drugs to elicit a selective effect at a remote site of action, liposomal impermeability should be maintained until arrival at the target site. Selective liposomal unloading requires great sophistication in design, but this is not needed when using local anesthetics, which are directly administered close to their local site of action. Thus, this specific limitation does not apply either to the existing commercial liposomal bupivacaine formulation or to the proliposomal ropivacaine studied by us.
3. Controlled Release at the Site of Action
Drug release at the site of action should be predictable and at a rate appropriate for clinical requirements. In our study, anesthetic effects were observed for at least 30 hours with high tissue drug levels at 4 days; similar pharmacokinetic–pharmacodynamic findings were also observed in volunteers.7 Although this rate of drug release is appropriate for clinical needs, it should be stated that the physicochemical properties that control drug unloading in proliposomal ropivacaine oil have not been investigated in this study and are not yet understood. Local drug release of liposomal local anesthetics has been reported to be unpredictable, may be affected by local temperature and pH, and may occur with an initial burst release17; these studies should also be repeated for proliposomal ropivacaine.
Local anesthetics are used for both perineural or neuraxial administration and local infiltration. There are only a few reports of the efficacy of perineural or neuraxial liposomal local anesthetics,18–20 and these do not currently meet consensus criteria for the use of investigational neuraxial drugs.21 By contrast, the clinical utility of liposomal local anesthetic wound infiltration has been demonstrated in peripheral surgery (bunionectomy,22 hemorrhoidectomy,23 breast augmentation,24 and total knee replacement25). Although hitherto, only bunionectomy and hemorrhoidectomy were Food and Drug Administration–approved applications for the commercially available liposomal bupivacaine, this limitation was revised in December 2015.b In these clinical scenarios, liposomal local anesthetics have been used to prolong the analgesic effect of local anesthetics without increasing complications or side effects.26,27
Finally, the lack of any adverse effects of the study drug on wound healing was tested in a porcine wound-healing model in this study. The porcine wound healing model has been reported to be the ideal animal model for cutaneous wound healing studies, with similar structure to human skin and a similar pattern of healing.28 Although the safe clinical application of this formulation to human surgical wounds cannot be assessed in volunteers for obvious ethical reasons, it should be confirmed by clinical trials of wound healing in patients.
Proliposomal ropivacaine exerted prolonged anesthesia with delayed elimination, typical for liposomal local anesthetics. The advantages of this novel proliposomal ropivacaine oil, compared with preprepared liposomal local anesthetics, are its ease of preparation and its extended shelf-stability (>2 years) at room temperature. E
Name: Elyad M. Davidson, MD.
Contribution: This author helped design the study, conduct the study, and collect the data.
Attestation: Elyad M. Davidson attests to the integrity of the analysis reported in this manuscript. He attests to having approved the final manuscript.
Conflicts of Interest: Elyad M. Davidson together with Yehuda Ginosar received a research grant ($10,000) from Pain Reform Ltd., Tel Aviv, through Hadassit Inc., Hadassah Hebrew University Medical Center, which was used to support volunteer and investigator costs in the clinical study. He has no other relationship with Pain Reform Ltd.
Name: Simon Haroutounian, PhD.
Contribution: This author together with Leonid Kagan was responsible for pharmacokinetic modeling.
Attestation: Simon Haroutounian attests to having approved the final manuscript.
Conflicts of Interest: Simon Haroutounian declares no conflicts of interest.
Name: Leonid Kagan, PhD.
Contribution: This author together with Simon Haroutounian was responsible for pharmacokinetic analysis.
Attestation: Leonid Kagan attests to having approved the final manuscript.
Conflicts of Interest: Leonid Kagan declares no conflicts of interest.
Name: Michael Naveh, Dr. Med Vet.
Contribution: As COO of Painreform Ltd., this author was the initiator and sponsor for the overall execution of the study.
Attestation: Michael Naveh attests to having approved the final manuscript.
Conflicts of Interest: Michael Naveh is an employee of Painreform Ltd., who hold patent rights to proliposomal ropivacaine (PRF-110) and its proliposomal platform.
Name: Arnon Aharon, MD.
Contribution: This author contributed to the study design, was the monitor for data collection, and was the archival author responsible for maintaining the study records.
Attestation: Arnon Aharon attests to the integrity of the original data and is the archival author responsible for maintaining the study records; he attests to having approved the final manuscript.
Conflicts of Interest: Arnon Aharon was a clinical studies consultant to Painreform Ltd. at the time of the study who hold patent rights to proliposomal ropivacaine (PRF-110) and its proliposomal platform.
Name: Yehuda Ginosar, BSc, MBBS.
Contribution: This author assisted with study design, performed the statistical analysis of the pharmacodynamic data, and drafted the final manuscript.
Attestation: Yehuda Ginosar attests to the integrity of the analysis reported in this manuscript.
Conflicts of Interest: Together with Elyad M. Davidson, Yehuda Ginosar received a research grant ($10,000) from Painreform Ltd., through Hadassit Inc., Hadassah Hebrew University Medical Center, which was used to support volunteer and investigator costs. He has no other relationship with Painreform Ltd.
This manuscript was handled by: Terese T. Horlocker, MD.
Plasma ropivacaine concentrations were determined by Nextar Chempharma Ltd., Ness Ziona, Israel. Cryo-TEM was performed at Ilse Katz Institute for Nanoscale Science and Technology, Ben Gurion University of the Negev, Beersheva, Israel.
1. Reddy KR. Controlled-release, pegylation, liposomal formulations: new mechanisms in the delivery of injectable drugs. Ann Pharmacother. 2000;34:915–23.
2. Grant SA. The Holy Grail: long-acting local anaesthetics and liposomes. Best Pract Res Clin Anaesthesiol. 2002;16:345–52.
3. Grant GJ, Lax J, Susser L, Zakowski M, Weissman TE, Turndorf H. Wound infiltration with liposomal bupivacaine prolongs analgesia in rats. Acta Anaesthesiol Scand. 1997;41:204–7.
4. Davidson EM, Barenholz Y, Cohen R, Haroutounian S, Kagan L, Ginosar Y. High-dose bupivacaine remotely loaded into multivesicular liposomes demonstrates slow drug release without systemic toxic plasma concentrations after subcutaneous administration in humans. Anesth Analg. 2010;110:1018–23.
5. Lian T, Ho RJ. Trends and developments in liposome drug delivery systems. J Pharm Sci. 2001;90:667–80.
6. Cohen R, Kanaan H, Grant GJ, Barenholz Y. Prolonged analgesia from Bupisome and Bupigel formulations: from design and fabrication to improved stability. J Control Release. 2012;160:346–52.
7. Ginosar Y, Haroutounian S, Kagan L, Naveh M, Aharon A, Davidson EM. Proliposomal ropivacaine oil: pharmacokinetic and pharmacodynamic data after subcutaneous administration in volunteers. Anesth Analg. 2016;122:1673–80.
8. Chaplan SR, Bach FW, Pogrel JW, Chung JM, Yaksh TL. Quantitative assessment of tactile allodynia in the rat paw. J Neurosci Methods. 1994;53:55–63.
9. Knudsen K, Beckman Suurküla M, Blomberg S, Sjövall J, Edvardsson N. Central nervous and cardiovascular effects of i.v. infusions of ropivacaine, bupivacaine and placebo in volunteers. Br J Anaesth. 1997;78:507–14.
10. Grant GJ, Barenholz Y, Bolotin EM, Bansinath M, Turndorf H, Piskoun B, Davidson EM. A novel liposomal bupivacaine formulation to produce ultralong-acting analgesia. Anesthesiology. 2004;101:133–7.
11. Betton D, Greib N, Schlotterbeck H, Joshi GP, Ubeaud-Sequier G, Diemunsch P. The pharmacokinetics of ropivacaine after intraperitoneal administration: instillation versus nebulization. Anesth Analg. 2010;111:1140–5.
12. Lefrant JY, de La Coussaye JE, Ripart J, Muller L, Lalourcey L, Peray PA, Mazoit X, Sassine A, Eledjam JJ. The comparative electrophysiologic and hemodynamic effects of a large dose of ropivacaine and bupivacaine in anesthetized and ventilated piglets. Anesth Analg. 2001;93:1598–605.
13. Tsibiribi P, Bui-Xuan C, Bui-Xuan B, Tabib A, Descotes J, Chevalier P, Gagnieu MC, Belkhiria M, Timour Q. The effects of ropivacaine at clinically relevant doses on myocardial ischemia in pigs. J Anesth. 2006;20:341–3.
14. Danielsson BR, Danielson MK, Böö EL, Arvidsson T, Halldin MM. Toxicity of bupivacaine and ropivacaine in relation to free plasma concentrations in pregnant rats: a comparative study. Pharmacol Toxicol. 1997;81:90–6.
15. Sokolsky-Papkov M, Golovanevski L, Domb AJ, Weiniger CF. Prolonged local anesthetic action through slow release from poly (lactic acid co castor oil). Pharm Res. 2009;26:32–9.
16. Weiniger CF, Golovanevski L, Domb AJ, Ickowicz D. Extended release formulations for local anaesthetic agents. Anaesthesia. 2012;67:906–16.
17. Weiniger CF, Golovanevski M, Sokolsky-Papkov M, Domb AJ. Review of prolonged local anesthetic action. Expert Opin Drug Deliv. 2010;7:737–52.
18. Viscusi ER, Candiotti KA, Onel E, Morren M, Ludbrook GL. The pharmacokinetics and pharmacodynamics of liposome bupivacaine administered via a single epidural injection to healthy volunteers. Reg Anesth Pain Med. 2012;37:616–22.
19. Ilfeld BM, Malhotra N, Furnish TJ, Donohue MC, Madison SJ. Liposomal bupivacaine as a single-injection peripheral nerve block: a dose-response study. Anesth Analg. 2013;117:1248–56.
20. Ilfeld BM. Liposome bupivacaine in peripheral nerve blocks and epidural injections to manage postoperative pain. Expert Opin Pharmacother. 2013;14:2421–31.
21. Eisenach JC, Shafer SL, Yaksh T. The need for a journal policy on intrathecal, epidural, and perineural administration of non-approved drugs. Pain. 2010;149:417–9.
22. Golf M, Daniels SE, Onel E. A phase 3, randomized, placebo-controlled trial of DepoFoam® bupivacaine (extended-release bupivacaine local analgesic) in bunionectomy. Adv Ther. 2011;28:776–88.
23. Gorfine SR, Onel E, Patou G, Krivokapic ZV. Bupivacaine extended-release liposome injection for prolonged postsurgical analgesia in patients undergoing hemorrhoidectomy: a multicenter, randomized, double-blind, placebo-controlled trial. Dis Colon Rectum. 2011;54:1552–9.
24. Minkowitz HS, Onel E, Patronella CK, Smoot JD. A two-year observational study assessing the safety of DepoFoam bupivacaine after augmentation mammaplasty. Aesthet Surg J. 2012;32:186–93.
25. Bramlett K, Onel E, Viscusi ER, Jones K. A randomized, double-blind, dose-ranging study comparing wound infiltration of DepoFoam bupivacaine, an extended-release liposomal bupivacaine, to bupivacaine HCl for postsurgical analgesia in total knee arthroplasty. Knee. 2012;19:530–6.
26. Viscusi ER, Sinatra R, Onel E, Ramamoorthy SL. The safety of liposome bupivacaine, a novel local analgesic formulation. Clin J Pain. 2014;30:102–10.
27. Portillo J, Kamar N, Melibary S, Quevedo E, Bergese S. Safety of liposome extended-release bupivacaine for postoperative pain control. Front Pharmacol. 2014;5:90.
28. Ansell DM, Holden KA, Hardman MJ. Animal models of wound repair: are they cutting it? Exp Dermatol. 2012;21:581–5.
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