# Repurposing chlorpromazine for anti-leukaemic therapy with the drug-in-cyclodextrin-in-liposome nanocarrier platform

Edvin Tang Gundersen, Zhiqiang Wang, Jan-Lukas Førde, Eric Larquet, Reidun Æsøy, Hugo Roussel, Lou Tosi, Gillian Barratt, Lars Herfindal, François-Xavier Legrand

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Edvin Tang Gundersen, Zhiqiang Wang, Jan-Lukas Førde, Eric Larquet, Reidun Æsøy, et al. Repurposing chlorpromazine for anti-leukaemic therapy with the drug-in-cyclodextrin-in-liposome nanocarrier platform. Carbohydrate Polymers, 2025, 358, pp.123478. 10.1016/j.carbpol.2025.123478. hal-04981168

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# Repurposing chlorpromazine for anti-leukaemic therapy with the drug-in-cyclodextrin-in-liposome nanocarrier platform☆

Edvin Tang Gundersen<sup>a,b,1</sup>, Zhiqiang Wang<sup>c,1</sup>, Jan-Lukas Førde<sup>a,d</sup>, Eric Larquet<sup>e</sup>, Reidun Æsøy<sup>a</sup>, Hugo Roussel<sup>c</sup>, Lou Tosi<sup>c</sup>, Gillian Barratt<sup>c</sup>, Lars Herfindal<sup>a,*,2</sup>, François-Xavier Legrand<sup>c,*,2</sup>

## ARTICLE INFO

**Keywords:**

Acute myeloid leukaemia
Chlorpromazine
Drug repurposing
Liposomes
Cyclodextrin
Zebrafish larvae

## ABSTRACT

Acute myeloid leukaemia (AML) accounts for 30 % of adult leukaemia cases, predominantly affecting individuals over 60. The standard “7 + 3” intensive chemotherapy regimen is unsuitable for many elderly patients, contributing to AML’s poor prognosis. While progress in drug therapies has been made, breakthroughs remain limited, indication-specific, and slow to expand. Drug repurposing offers a faster route to therapy development, while nanocarrier encapsulation broadens the scope of viable drug candidates.

Chlorpromazine (CPZ) is an antipsychotic which has been identified as a potential anti-leukaemic agent. Due to its ability to cross the blood-brain barrier, it is likely to cause central nervous system (CNS) effects. The drug-in-cyclodextrin-in-liposome (DCL) nanocarrier platform enables the formulation of CPZ encapsulated with cyclodextrins (CDs) such as HP-γ-CD, SBE-β-CD, and Sugammadex.

The CD/CPZ formulations were equally, or more efficient than free CPZ in inducing AML cell death. Uptake of the DCL in AML cells quickly reached saturation, with minimal differences among formulations, except for SBE-β-CD. When injected intravenously in zebrafish larvae, the different DCLs did not differ in biodistribution, and no brain accumulation was observed at two days post-injection. These DCL-based CPZ formulations maintain anti-leukaemic activity, avoid CNS accumulation, and allow drug availability adjustments based on the included CD.

## 1. Introduction

Acute myeloid leukaemia (AML) accounts for 30 % of all leukaemia in adults and has a poor prognosis, particularly in older patients (Rodriguez-Abreu et al., 2007). The most common treatment regimen for AML, the standard “7 + 3 regimen” (cytarabine for 7 days with daunorubicin or idarubicin for 3 days) comes with severe side effects as well as high relapse rates. There is a need for new treatments, and several small molecules have recently been launched or are under investigation, including several multi-kinase inhibitors like clofarabine, sorafenib, midostaurin and crenolanib (Montalban-Bravo & Garcia-

Manero, 2015). New treatments can also be developed by drug repurposing, a strategy that directs well-established drugs and molecules towards different diseases, and thus offers the possibility of reducing the time, cost, and risk associated with conventional drug discovery and development (Andresen & Gjertsen, 2017; Ashburn & Thor, 2004; Gupta et al., 2013). In this respect, chlorpromazine (CPZ), a widely used antipsychotic, was reported to limit the growth of murine leukaemic lymphocytes (L1210 and L5178Y) and human promyelocytic leukaemic granulocytes (HL-60) (Hait et al., 1985). CPZ showed higher cytotoxicity against leukaemic cell lines than its sulphoxide derivative, with IC₅₀ values of 8 µM and 100 µM respectively on the HL-60 cell line. CPZ has

<footnote><sup>a</sup> Centre for Pharmacy, Department of Clinical Science, University of Bergen, Bergen, Norway</footnote>

<footnote><sup>b</sup> Hospital Pharmacies Enterprise, Western Norway, Bergen, Norway</footnote>

<footnote><sup>c</sup> Université Paris-Saclay, CNRS, Institut Galien Paris-Saclay, 91400 Orsay, France</footnote>

<footnote><sup>d</sup> Department of Internal Medicine, Haukeland University Hospital, Bergen, Norway</footnote>

<footnote><sup>e</sup> Laboratoire de Physique de la Matière Condensée (PMC), CNRS, Ecole Polytechnique, Institut Polytechnique de Paris, 91120 Palaiseau, France</footnote>

<footnote>* This article is part of a special issue entitled: Cyclodextrin conference published in Carbohydrate Polymers.</footnote>

<footnote>* Corresponding authors.</footnote>

<footnote>E-mail addresses: lars.herfindal@uib.no (L. Herfindal), francois-xavier.legrand@universite-paris-saclay.fr (F.-X. Legrand).</footnote>

<footnote n="1">¹ These authors contributed equally.</footnote>

<footnote n="2">² These authors contributed equally.</footnote>
therefore been identified as a candidate drug to be repurposed to AML therapy (Sachlos et al., 2012).

As an anti-psychotic drug, CPZ passes through the blood-brain barrier (BBB) and accumulates in the central nervous system (CNS). This is problematic in non-psychotic individuals, where CPZ causes a series of extrapyramidal side effects such as involuntary shaking and movements. Therefore, a new strategy is needed for repurposing CPZ, one which prevents it from accumulating in the brain while delivering it to the AML target cells. The use of nanosized drug carriers is a powerful method of modulating drug distribution and pharmacokinetics without altering the drug itself. Liposomes are the earliest example of nano-metric drug delivery systems and have been widely used over the past 50 years since their first discovery in 1965 (Bangham et al., 1965). Numerous agents, including hydrophobic, hydrophilic, and amphiphilic molecules can be encapsulated in the lipid vesicles and transported through the circulatory system after administration. A major step forward in liposome design came with the development of so-called "Stealth"-systems that avoid opsonisation and uptake by phagocytic cells and can therefore persist in the circulation and deliver drugs to sites such as solid tumours. A number of liposomal systems have been commercialised after FDA approval such as Doxil®/Caelyx®, a PEGylated liposomal formulation of doxorubicin indicated for Kaposi's sarcoma, ovarian cancer, and multiple myeloma (Olusanya et al., 2018), and Vyxeos®, a liposomal formulation co-encapsulating daunorubicin and cytarabine for use against newly diagnosed therapy-related AML (t-AML) and AML with myelodysplasia-related changes (AML-MRC) (Briot et al., 2018).

It was hypothesised that encapsulating CPZ in liposomes could effectively prevent its passage through the BBB, extend its plasma circulation time, and enhance its accumulation and uptake in the bone marrow. Moreover, the liposomal formulation provides the possibility of active targeting for the drug, wherein the liposome surface is decorated with appropriate monoclonal antibodies or fragments or other ligands that can recognise surface epitopes on AML cells and preferentially associate. However, considering the amphiphilic nature CPZ and the possibility of its interaction with the phospholipid bilayer, rapid release may occur before the liposomes have reached their target in the bone marrow. To avoid premature release of CPZ, the drug-in-cyclodextrin-in-liposomes (DCL) strategy was adopted, wherein CPZ is encapsulated in liposomes as a complex with a cyclodextrin (CD). This concept was first proposed by McCormack and Gregoriadis in 1994 (McCormack & Gregoriadis, 1994; McCormack & Gregoriadis, 1996), who created CD/drug-complexes with various hydrophobic drugs, which they then encapsulated in the aqueous compartment of liposomes to control release rate and biodistribution in rats. More recently, Gallez et al. observed differences in activity for 17β-estradiol administered as the free drug and as DCL; although both forms allowed the genomic, nuclear activity of the drug, the DCL form did not induce non-genomic, extranuclear effects that were mediated by cell-membrane receptors, demonstrating that encapsulation could modify the distribution (Gallez et al., 2020).

A wide range of CDs has been previously studied for their ability to form high-affinity complexes with CPZ (Wang et al., 2020), leading to the selection of three CDs for further investigation (Table 1): hydroxypropyl-γ-cyclodextrin (HP-γ-CD), sulphobutylether-β-cyclodextrin (SBE-β-CD), and the γ-cyclodextrin Sugammadex (SGM). While the latter two showed the highest complexation ratios due to higher affinity for CPZ, HP-γ-CD was chosen because this CD has the lowest complexation ratio towards CPZ among the CDs that are well accepted as pharmaceutical excipients (Table 1) (Wang et al., 2020). Furthermore, these three CDs were also chosen as they do not disrupt the structure of liposomes (Wang, 2019). The data presented herein describe the formulation and characterisation of liposomes containing CPZ alone or CPZ in complexation with one of three CDs. An evaluation of the toxicity of these formulations towards a panel of leukaemic cell lines, as well as a non-cancerous cell line, was then performed. Liposomes with or without

<table><caption>**Table 1**
Structural details of cyclodextrins used in this study and their complexation with chlorpromazine.</caption><thead><tr><th>General structure</th><th>Abbreviation</th><th>n</th><th>R<sub>2</sub></th><th>R<sub>3</sub></th><th>R<sub>6</sub></th><th>DS</th><th>Association Constant</th><th>Complexed CPZ (%)</th><th>Total complexed CPZ (%)</th></tr></thead><tbody><tr><td rowspan="4"></td><td>HP-γ-CD</td><td>8</td><td></td><td>OCH<sub>2</sub>CH(OH)CH<sub>3</sub> or OH</td><td></td><td>6.6</td><td>K<sub>1,1</sub> = 4.57 × 10<sup>-2</sup> M<sup>-1</sup></td><td>23.6</td><td>74.7</td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td>K<sub>1,2</sub> = 1.07 × 10<sup>-3</sup> M<sup>-1</sup></td><td>51.1</td><td></td></tr><tr><td>SBE-β-CD</td><td>7</td><td></td><td>OCH<sub>2</sub>CH<sub>2</sub>CH<sub>2</sub>SO<sub>3</sub>Na or OH</td><td></td><td>5.8</td><td>K<sub>1,1</sub> = 1.99 × 10<sup>-4</sup> M<sup>-1</sup></td><td>60.6</td><td>96.5</td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td>K<sub>1,2</sub> = 2.09 × 10<sup>-3</sup> M<sup>-1</sup></td><td>35.9</td><td></td></tr><tr><td></td><td>SGM</td><td>8</td><td>OH</td><td>OH</td><td>SCH<sub>2</sub>CH<sub>2</sub>CO<sub>2</sub>Na</td><td>8</td><td>K<sub>1,2</sub> = 6.37 × 10<sup>-9</sup> M<sup>-2</sup></td><td>97.9</td><td>97.9</td></tr></tbody></table>

HP-γ-CD: hydroxypropyl-γ-cyclodextrin, SBE-β-CD: sulphobutylether-β-cyclodextrin, SGM: Sugammadex, DS: degree of substitution.

Association constants at 25 °C are taken from (Wang et al., 2020). The percentage of complexed and total complexed CPZ are calculated at 25 °C for a CPZ concentration of 4 mM and a CD concentration of 4 mM.
CDs and modified with a fluorescent phospholipid were also investi-
gated for cellular uptake, as well as biodistribution after intravenous
injection into zebrafish larvae. This model was chosen because the
zebrafish BBB closely resembles that of mammals in structure and
function, shares significant sequence similarity in transporters and re-
ceptors with humans, matures approximately 2.5 days post-fertilisation,
allows real-time pharmacokinetic tracking due to larval transparency,
and enables rapid and reliable BBB permeability assessment alongside
CNS-targeted behavioral assays comparable to those used in rodent
models (Kim et al., 2017; Quiñonez-Silvero et al., 2020; Stanimirovic
et al., 2015).

## 2. Materials and methods

### 2.1. Materials

Hydroxypropyl-γ-cyclodextrin (HP-γ-CD, DS = 5.8, Cavasol® W8HP Pharma) was a gift from Ashland Global Specialty Chemicals Inc., sulphobutylether-β-cyclodextrin (SBE-β-CD, DS = 6.6, Captisol®) was a gift from Ligand Pharmaceuticals Inc., and Sugammadex (SGM, γ-cyclodextrin selectively persubstituted on the primary face with 3-mercapto-propionic acid, Bridion®) was obtained from Bichat Hospital.

Hydrogenated egg L-α-phosphatidylcholine (HEPC), cholesterol, 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] ammonium salt (DSPE-PEG<sub>2000</sub>), and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulphonyl) ammonium salt (18:1 Liss Rhod PE), were all supplied by Avanti Polar Lipids Inc. (Alabaster, AL, USA).

Chlorpromazine hydrochloride (CPZ), 4-(2-hydroxyethyl)-1-piperazineethanesulphonic acid (HEPES), phosphate-buffered saline (PBS), Tween® 20, sodium chloride, formaldehyde, octyl glucoside, tricaine (ethyl 3-aminobenzoate methane sulphonate), ammonium sulphate, ammonium bicarbonate, cell culture media (RPMI 1640, IMDM, DMEM), foetal bovine serum (FBS), L-glutamine, penicillin, and streptomycin were purchased from Sigma-Aldrich (St. Louis, MO, USA). ProLong® Gold Antifade Mounting medium with DAPI, Hoechst 33342 fluorescent DNA staining reagent, and FBS was purchased from Thermo Fisher Scientific (Waltham, MA, USA). WST-1 cell proliferation reagent was purchased from Roche Diagnostics GmbH (Mannheim, Germany). Milli-Q water was obtained by a Millipore® purification system (Millipore, Germany).

E3 medium with methylene blue was provided by the Zebrafish facility at the Department of Bioscience, University of Bergen.

Calcium chloride was purchased from Merck (Darmstadt, Germany), while calcium lactate was purchased from Fisher Scientific (Illkirch, France). Diethylmalonylurea sodium salt and diethylmalonylurea acid were purchased from Prolabo (Paris, France). Magnesium chloride, tricine, tris, and agarose were purchased from Sigma-Aldrich (Saint-Quentin-Fallavier, France). Coomassie brilliant blue was purchased from Thermo Fisher Scientific (Les Ulis, France). Polyclonal anti-human C3 antibodies raised in goat were purchased from Fitzgerald antibodies (Acton, MA, USA).

Human serum was prepared from human plasma provided by the
Établissement Français du Sang (Agreement 14/EFS/041, Rungis,
France).

### 2.2. Liposome preparation

The drug-in-cyclodextrin-in-liposome (DCL) formulations were pre-
pared using the dehydration-rehydration vesicle (DRV) method that was
first developed by Kirby and Gregoriadis (Kirby & Gregoriadis, 1984).
This method has been widely used to prepare liposomal formulations
containing various substances, in particular large hydrophilic molecules
such as proteins and nucleic acids, and has already been applied to
cyclodextrin (CD) complexes (Antimisiaris, 2017). It consists of first
preparing small unilamellar vesicles (SUV) and freeze-drying them the

presence of the substance to be encapsulated. Since preliminary exper-
iments with this method resulted in low encapsulation yields of chlor-
promazine (CPZ) or CD/CPZ complexes, an ammonium sulphate loading
procedure was added to the preparation process to increase the lipo-
somal encapsulation of CPZ both by itself and in its CD-complexed form.
Thus, the rehydration step after freeze-drying was performed using a
300 mM solution of ammonium sulphate that can form an insoluble
complex with both CPZ and ionisable CDs, favouring retention in the
liposomes during the subsequent purification steps (Gabizon et al.,
1989).

To prepare the initial SUV, multilamellar liposomes composed of HEPC, cholesterol, and DSPE-PEG<sub>2000</sub> (65:30:5, molar ratio) were first obtained by the film hydration method. The components were dissolved in a mixture of chloroform and methanol (2:1, v/v) and the organic solvents were evaporated at room temperature under a gentle stream of nitrogen, followed by drying under a vacuum overnight to completely remove any residual solvent. An appropriate volume of water was filtered by two passages through sterile filters (0.2 µm pore size) and added to the film to obtain a total lipid concentration of 36 mM. The lipid suspension subsequently underwent 10 repeated cycles of 1 min heating at 62 °C and 1 min of vortexing. SUVs were prepared by extrusion through polycarbonate membranes (1 passage through 0.4 µm pore size, 2 passages through 0.2 µm pore size, 10 passages through 0.1 µm pore size) at 62 °C using a LIPEX® liposome extruder (Northern Lipids, Burnaby, Canada).

DRV were then prepared by adding either 2 mL of a CPZ solution (4 mM CPZ in Milli-Q purified water), or 2 mL of an aqueous solution containing 4 mM CPZ and 4 mM CD (HP-γ-CD, SBE-β-CD, or SGM), to a 2 mL suspension of SUV with a total lipid concentration of 36 mM, followed by vigorous mixing with a vortex mixer for 3 min. The mixture was frozen at -20 °C for at least 3 h, followed by overnight lyophili-sation in a Cosmos freeze-dryer (Cryotec, Saint-Gély-du-Fesc, France). For rehydration, 0.2 mL of 300 mM ammonium sulphate solution was pre-heated to 62 °C and added to the freeze-dried lipid pellet, which was then heated at 62 °C and vortexed for several cycles until completely hydrated. After leaving the dispersion at room temperature for 30 min, 1.8 mL of HEPES buffer was added, with the mixture then kept at 62 °C for 50 min. Since this procedure led to an increase in liposome diameter, a second extrusion step was performed with two passages through a 0.2 µm polycarbonate membrane. Non-encapsulated CPZ by was removed by ultracentrifugation in a Beckman Optima L-90K ultracentrifuge (Beckman Coulter, Brea, CA, USA) equipped with a Type 70:1 Ti rotor at 146,682 × g at 4 °C for 6 h. The liposome pellet was resuspended in 1.5 mL HEPES buffer. These procedures are illustrated in Supplementary Fig. S1.

Fluorescent liposomes composed of HEPC, cholesterol, DSPE-PEG<sub>2000</sub>, and 18:1 Liss Rhod PE (65:30:5:0.5, molar ratio), without encapsulated CPZ, were prepared by the same method.

### 2.3. Size and zeta potential measurements

The mean hydrodynamic diameters of the liposome preparations were obtained by dynamic light scattering using a Zetasizer Nano ZS90 (Malvern Instruments Corp., Worcestershire, UK). For size determination, 14 µL of sample was diluted in 1 mL HEPES buffer and measurements were performed at 25 °C, with a detection angle of 90°. Size distributions by intensity were recorded and the polydispersity index (PDI) was also acquired to check the size distribution. Zeta potential, as an indication of the liposome surface charge, was evaluated using the same instrument by diluting 20 µL of sample in 1 mL of ultrapure water and measuring three independent aliquots of 1 mL at 25 °C. Data were recorded using the general-purpose mode and the Smoluchowski limit was used to convert electrophoretic mobility to zeta potential.
## 2.4. Phospholipid assay

The phospholipid concentrations of the liposome preparations were estimated by measuring phosphatidylcholine using a commercial kit (Phospholipids Colorimetric Enzymatic Method, BIOLABO, France) according to the instructions of the manufacturer, but with the modification of diluting the liposome suspensions in a solution of the detergent n-octyl-β-D-glucoside (final concentration of 200 mM) to disperse the lipids as micelles. The total lipid concentration was then calculated from the proportion of phosphatidylcholine in the preparation, assuming that all lipids were recovered with the same yield and the average lipid molecular weight was 776 Da.

## 2.5. Cryo-electron microscopy

Samples for cryo-electron microscopy were prepared by depositing 4 µL of suspension, with a lipid concentration around 36 ± 4 mg/mL for undiluted samples and around 12 ± 1 mg/mL for diluted samples, on a 200-mesh lacey carbon-coated grid. After blotting with filter paper, the grid was frozen by rapid plunging into liquid ethane, then mounted and inserted in the microscope using a nitrogen-cooled side entry Gatan Elsa™ (698) Cryo-holder at a temperature of -181 °C. Observations were carried out in a Thermo Fisher Scientific Titan Themis 300 transmission electron microscope equipped with a Cryo-box anti-contamination system and a C-Twin objective lens (Cs, 2.7 mm; Cc, 2.7 mm), using an accelerating voltage of 300 kV, with the following illumination conditions: spot size of 5, a 150 µm condenser aperture, and no objective aperture. Images were recorded with a defocus range of -1.2 to -2.0 µm, using the low electron dose system of 10 electrons per Å² per s, with a magnification of 28,500× and 58,000× on a Thermo Fisher Scientific Falcon 3 Direct Detection Electron 4 K × 4 K camera in linear mode.

## 2.6. Quantification of CPZ by HPLC

High Performance Liquid Chromatography (HPLC) analyses of CPZ concentrations were performed using a Waters 717 Plus Autosampler HPLC system equipped with Waters 2487 Dual λ Absorbance Detector at wavelength of 254 nm and 309 nm. A C18 chromatographic column (Waters XTerra®, 5 µm, 3.0 × 150 mm) thermostated at 30 °C was used for separation. The mobile phase was comprised of a mixture of acetonitrile, water, and 0.1 mM ammonium bicarbonate pH 7 (7:2:1, v/v/v). The flow rate was set at 0.7 mL/min, and the injection volume at 10 µL. A linear calibration curve was prepared using CPZ in mobile phase at concentrations between 5 µg/mL and 100 µg/mL.

## 2.7. Determination of the encapsulation efficiency and drug loading of CPZ

The encapsulation efficiency (EE) and drug loading (DL) of CPZ in liposomes were determined after separating encapsulated and non-encapsulated CPZ by ultracentrifugation, as described above, and mixing 0.1 mL of the dispersed DRV pellet with 0.4 mL HPLC mobile phase solution and performing HPLC as described above. The CPZ concentration was calculated from a calibration curve and the EE was obtained by Eq. (1), and together with the lipid concentration determined by the phospholipid assay, as described above, the DL was obtained by Eq. (2) (Sauvage et al., 2018).

$$EE (\%) = \frac{\text{CPZ concentration in DRV (mM)}}{\text{Initial CPZ concentration (mM)}} \times 100 \quad (1)$$

$$DL (\%) = \frac{\text{CPZ concentration in DRV (mM)}}{\text{Lipid concentration (mM)}} \times 100 \quad (2)$$

## 2.8. Cell lines and culture

Four human leukaemic cell lines were used in this study: the human acute promyelocytic leukaemia cell line HL-60 (ATCC CCL-240) provided by Dr. Brigitte Bauvois (INSERM U872, Paris), the human AML cell lines MOLM-13 (ACC 554) (Matsuo et al., 1997; Quentmeier et al., 2003) and OCI-AML3 (ACC 582), and the human monocytic leukaemia cell line MV4-11 (ATCC CRL-9591). The normal rat kidney epithelial cell line NRK (ATCC CRL-6509) was used as a non-cancerous cell line. HL-60, MOLM-13 and OCI-AML3 were cultured in RPMI 1640 medium, while MV4-11 cells were cultured in IMDM medium, and NRK cells in DMEM medium. All media were supplemented with 10% foetal bovine serum, 100 IU/mL penicillin, 0.1 mg/mL streptomycin, and L-glutamine to a final concentration of 20 mM. Cells were maintained in a humidified incubator at 37 °C with 5% CO₂. Mild trypsin treatment (0.33 mg/mL trypsin for 5 min at 37 °C) was used to detach NRK cells when they had reached 80% confluence, followed by centrifugation at 160 × g for 4 min and reseeding in fresh medium (Haji Ibrahim et al., 2019).

## 2.9. Cytotoxicity evaluation

The cytotoxicity of the different formulations was assessed by metabolic assay, using tetrazolium-based substrates that are reduced by mitochondrial enzymes leading to products that can be measured spectrophotometrically. In the case of HL-60, viability was evaluated by MTS metabolism. Briefly, 25,000 cells in 50 µL medium per well were seeded in 96-well plates and incubated for 4 h, after which 50 µL medium containing different concentration of CPZ, CDs, empty liposomes, liposomes containing CPZ, or liposomes containing CD/CPZ complexes, were added. After incubation for up to 72 h, cell viability was assessed by CellTiter 96® Aqueous One Solution Cell Proliferation Assay (Promega Corporation, Madison, WI, USA), wherein 25 µL of MTS reagent was added to each well and the plates were incubated for a further 2 or 4 h, after which the optical density was measured with a high-pass filter at 490 nm with an LT-5000 ELISA plate reader (Labtech, Heathfield, UK).

The cytotoxicity towards MOLM-13, OCI-AML3, and MV4-11 was assessed by a similar method measuring the conversion of WST-1. Briefly, 50 µL medium containing different concentrations of drugs was first prepared in the wells of 96-well plate, followed by adding 50 µL cells in medium to each well (final cell count per well: 35,000 for 24-hour incubation and 10,000 for 72-hour incubation). To assess metabolic activity of the cells, 10 µL of WST-1 reagent was added to each well and the plate was incubated for another 2 h. The optical density was then measured with a TECAN Infinite M200 PRO microplate reader with the Magellan software version 7.2 (Tecan Group Ltd., Männedorf, Zürich, Switzerland). The absorbance and reference wavelengths were 450 nm and 620 nm, respectively.

When testing on NRK cells, 96-well plates were prepared by seeding the cells in 100 µL medium per well and incubated overnight to attach. The number of cells seeded per well was 5000 and 1000 for 24- and 72-hour experiments, respectively. The medium was then removed and fresh medium of equal volume containing different concentrations of drugs was added, and the cells were incubated for 24 or 72 h before assessment of viability by WST-1.

In all relevant experiments, wells containing the different dilutions of the formulations, but not any cells, were measured and used as blanks to correct for possible interference caused by the absorbance of the formulations.

Cell death kinetics experiments were performed on MOLM-13 cells seeded in 96-well plates with 35,000 cells per well. Cells were incubated up to 12 h with 100 µM of either free or encapsulated CPZ, added to wells at given time points. Empty DRV of equivalent lipid concentration to the most concentrated CPZ formulation were also tested. After incubation, cells were fixed in 2% formaldehyde fix in pH 7.4 PBS, stained with 0.01 mg/mL Hoechst 33342, and counted for apoptotic nuclei with
an Axiovert 200M fluorescence microscope (Carl Zeiss AG, Oberkochen, Germany) to determine cell death. Image analysis and processing was done using the ImageJ software version 1.52p (U. S. National Institute of Health, Bethesda, MD, USA) (Schneider et al., 2012), as a part of the FIJI image processing package (Schindelin et al., 2012).

## 2.10. Flow cytometry

To investigate potential time-dependent cellular uptake or cell surface association of liposomes, MOLM-13 cells were incubated for up to 8 h with 2 % v/v concentration of rhodamine-labelled fluorescent dehydration-rehydration vesicles (F-DRV) made either with or without CDs. Cells were seeded in wells of a 48-well plates at 350,000 cells/mL and incubated in growth conditions (37 °C with 5 % CO₂). After incubation, the cells were washed twice with cold PBS by centrifugation at 300 × g for 10 min to remove supernatant, and then kept dark and on ice until flow cytometric analysis. The fluorescence intensity of the cells was measured with a FACS Accuri C6 flow cytometer and analysed using the Accuri CFlow software version 1.0.227.4 (BD Biosciences, San Jose, CA, US). The following instrument settings were used: 488 nm laser, 585/40 nm filter, and fast fluidics (66 µL/min). At least 10,000 events were recorded when gated for living cells, with further gating done to exclude cell doublets and aggregates (see Supplementary Fig. S2 for gating examples). Steady-state fluorescence measurements were performed using a SPEX Fluorolog spectrofluorimeter at 25 °C with a Liss Rhod PE concentration of approximately $10^{-8}$ mol.L⁻¹. Emission spectra were recorded from 578 to 700 nm with an excitation wavelength of 568 nm, while excitation spectra were recorded from 450 to 578 nm with an emission wavelength of 588 nm (Supplementary Fig. S3). To account for differences in fluorescence output between F-DRV formulations, dilution series (0.16–10 %, v/v) of the formulations were made in PBS with 5 % Tween® 20 and measured for fluorescence with a PerkinElmer EnVision 2103 Multilabel plate reader with version 1.12 of the Wallac EnVision Manager software (PerkinElmer, Waltham, MA, USA) (Supplementary Fig. S4). Excitation and emission wavelength were set to 485 and 590 nm.

## 2.11. Confocal microscopy

Cell uptake or association of fluorescent liposomes was also investigated by confocal microscopy. MOLM-13 cells were treated and washed as described for flow cytometry but after wash steps were resuspended in 2 % formaldehyde fixative and kept in the dark for at least 30 min at room temperature before replacing the fixative with PBS. The cells were spun down onto glass slides at 300 × g for 10 min using a Shandon Cytospin 3 centrifuge and mounted using ProLong® Gold mounting medium with DAPI. Confocal images were obtained using an Andor Dragonfly 505 confocal microscope (Andor Technologies Inc., Belfast, UK) fitted with a CFI SR HP Apo TIRF 100× objective. A 488 nm Argon laser was used to excite the rhodamine fluorochrome, while a 405 nm UV laser was used visualise DAPI-stained nuclei.

## 2.12. Zebrafish maintenance, injection, and imaging

To examine DRV biodistribution, fluorescent rhodamine-labelled F-DRV were injected into the larvae of the zebrafish (Danio rerio) animal model (Santoriello & Zon, 2012), specifically of the optically transparent casper line, seen as an ideal model system for microscopy-based screening of anti-cancer nanocarriers (D'Agati et al., 2017; Evensen et al., 2016; White et al., 2008). Zebrafish embryos were obtained from the Zebrafish Facility at the Department of Biological Sciences, University of Bergen. The facility operates in accordance with the European Convention for the Protection of Vertebrate Animals Used for Experimental and Other Scientific Purposes. Fertilized eggs were collected, and larvae were used prior to the development of sex dimorphism, making it impossible to distinguish between sexes. The embryos were maintained

in fresh E3 medium at 28.5 °C and anesthetized in a 0.7 mM tricaine solution before and during injection and imaging. At around two days post-fertilisation (dpf), i.e. the long-pec stage (48 h) of the hatching period (48–72 h) as described by Kimmel et al. (Kimmel et al., 1995), hatched or dechorionated zebrafish larvae were intravenously injected with approximately 4 nL of rhodamine-labelled F-DRV via the posterior cardinal vein. The larvae were imaged on the day of injection, within three hours post-injection (hpi), i.e. zero days post-injection (dpi), and next at one- and two-days post-injection. Imaging was done with an Andor Dragonfly 505 confocal microscope fitted with a CFI Plan Apochromat Lambda 10× objective, 561 nm excitation laser, and 600/25 nm band pass filter. Fluorescence images of larvae are a Z-projection obtained by summation of overlying pixels in the confocal Z-stack, while 3D representations of the blood vessels in the brain were obtained by pre-processing using a 3D-median filter. Image analysis, processing, and rendering was done using the ImageJ software version 1.52p as a part of the FIJI image processing package (Schindelin et al., 2012; Schneider et al., 2012).

## 2.13. Complement activation

The ability of the different liposome formulations to activate the complement system was assessed using a miniaturised crossed immunoelectrophoresis system as described by Coty et al. (Coty et al., 2016). Briefly, human serum (25 µL) was incubated with 50 µL liposome suspension adjusted in concentration to contain the same particle surface area of 4000 cm²/mL for each formulation, and 25 µL of Veronal-buffered saline (VBS²⁺) containing both Ca²⁺ and Mg²⁺ ions (1.5 mM diethylmalonylurea sodium salt, 3 mM diethylmalonylurea acid, 0.015 mM calcium chloride and 0.05 mM magnesium chloride, pH 7.5). Surface area was calculated from the lipid concentration and the mean hydrodynamic radius, assuming that the density of the liposomes was 1 g/cm³. The negative control was HEPES buffer, and the positive control, designated as A1, was poly(isobutylcyanoacrylate) nanoparticles prepared by anionic emulsion polymerisation with side-on dextran chains at the surface as described in Coty et al. (Coty et al., 2017), known to be a strong complement activator and adjusted to the same total surface area as the liposomes. The samples were incubated for 1 h at 37 °C and immediately cooled to 4 °C in an ice-bath. An electrophoretic gel was prepared from 1 % agarose in Tricine buffer (27 mM Tricine, 63 mM Tris and 1 mM calcium lactate, pH 8.5). The gel was divided in 35 subsections of 2.5 cm by 2.5 cm. Each sub-section contained a band 8 mm wide containing plain agarose and a sample well, while the rest of the gel was cast from agarose containing polyclonal anti-human C3 antibodies raised in goats. Electrophoresis was first run along one dimension in the plain section of the gel to separate serum proteins by molecular size, then along a second dimension in the antibody-loaded section of the gel to identify intact and cleaved C3 by rocket immunoelectrophoresis. The gel was then dried and stained with Coomassie brilliant blue. After scanning the gel, the areas of the peaks corresponding to native C3 and C3 fragments were estimated using ImageJ. The percentage of activation was expressed as the ratio of the peak area of C3 fragments to that of the sum of the peaks of intact C3 and its fragments. Two or three replicate gels were run for each liposome preparation or control. Spontaneous activation in the serum was checked by loading serum incubated with Veronal-buffered saline alone.

# 3. Results and discussion

## 3.1. Liposome preparation and characterisation

The dehydration-rehydration method of preparing loaded liposomes was chosen for the encapsulation of cyclodextrin/chlorpromazine (CD/CPZ) complexes because it was reported to give high encapsulation efficiency for high molecular weight water-soluble molecules. This approach has already been used by several groups to encapsulate CD/
<table><caption>**Table 2**
Properties of the liposomal formulations used in this work.</caption><thead><tr><th>Liposome formulation</th><th>Final mean diameter (nm)</th><th>PdI</th><th>Zeta Potential (mV)</th><th>Lipid conc. (mM)</th><th>CPZ conc. (mM)</th><th>CPZ EE (%)</th><th>DL (%)</th></tr></thead><tbody><tr><td>Empty DRV</td><td>197 ± 7</td><td>0.080</td><td>-30.0 ± 5.7</td><td>19.6</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>DRV + CPZ</td><td>183 ± 1</td><td>0.056</td><td>-15.6 ± 6.2</td><td>23.6</td><td>1.31</td><td>32.8</td><td>5.6</td></tr><tr><td>DRV + HP-γ-CD/CPZ</td><td>177 ± 6</td><td>0.107</td><td>-19.4 ± 5.9</td><td>23.1</td><td>0.83</td><td>20.8</td><td>3.6</td></tr><tr><td>DRV + SBE-β-CD/CPZ</td><td>187 ± 3</td><td>0.070</td><td>-24.7 ± 5.2</td><td>23.1</td><td>0.66</td><td>16.5</td><td>2.9</td></tr><tr><td>DRV + SGM/CPZ</td><td>186 ± 6</td><td>0.063</td><td>-28.2 ± 5.4</td><td>24.9</td><td>0.48</td><td>12.0</td><td>1.9</td></tr></tbody></table>

DL: Drug loading, EE: Encapsulation efficiency, PdI: Polydispersity index.

drug complexes, for example betamethasone (Piel et al., 2006), keto- profen (Maestrelli et al., 2005), and curcumin (Dhule et al., 2012). To further improve the encapsulation, the ammonium sulphate trapping method was included to allow for CPZ, which is a weak base, to form an insoluble complex, thus limiting its leakage into the aqueous phase. With this method, the EE obtained for CPZ by itself was just above 30 %, as shown in Table 2. However, when liposome formulations including CDs were prepared by the same method, the EE of CPZ obtained was only between 12 and 21 % (Table 2). This could be because a high proportion of the CPZ would be expected to be included as a complex with the chosen CD, and thus not be subject to ammonium sulphate trapping. The observed encapsulation rate therefore probably reflects that of the chosen CD. In the literature, differences in drug encapsulation have been observed between drug and CD-associated drug, and between different CDs. According to the study of Piel et al., the encapsulation efficiency of betamethasone showed marked differences between CD types (Piel et al., 2006). The dependence of drug EE on the CD con- centration in a DCL system was investigated by Maestrelli et al., who reported that the EE of ketoprofen reached a maximum when increasing of HP-β-CD/ketoprofen complex concentration to 10 mM CD, then declined. This decrease of EE was explained by the destabilising effect of CD at high concentration (Maestrelli et al., 2006). However, in our ex- periments, the CD concentration used was only 4 mM. In a review by Gharib et al. it is pointed out that the use of HP-β-CD often leads to higher encapsulation efficiency than for drug without CD because of the high aqueous solubility of this particular CD (Gharib et al., 2015).

Table 2 also shows the mean hydrodynamic diameters of represen- tative liposome preparations. After the freeze-drying step, the recon- stituted liposomes were found to be larger and more polydisperse than the original small and unilamellar liposomes. This was expected because it is difficult to preserve liposome structure during freeze-drying unless a specific cryoprotective agent is added (Crowe et al., 1985). Therefore, a second extrusion step with a 0.2 µm membrane was performed to obtain formulations suitable for intravenous injection. The subsequent ultra- centrifugation procedure to separate free and encapsulated CPZ did not affect the liposome size. Typical dynamic light scattering (DLS) profiles of rehydrated, re-extruded, and final liposome preparations after ultra- centrifugation, are given in Supplementary Fig. S5. The final liposomes had diameters close to 200 nm, and polydispersity indices of 0.1 or lower, except for HP-γ-CD/CPZ liposomes which had a higher poly- dispersity index with multiple populations sometimes observed

(Supplementary Fig. S5C and Table 2).

Cryo-electron microscopy (Cryo-EM) gave complementary informa- tion to the DLS results (Fig. 1). All the preparations showed mainly unilamellar vesicles with diameters between 100 and 200 nm and no chlorpromazine nanocrystals or fibers are observed in the loaded for- mulations, unlike the classical observations with anthracyclines (Reiten et al., 2025). However, the HP-γ-CD/CPZ liposomes showed a wider size range (Fig. 1C) in accordance with the higher polydispersity index. In addition to spherical vesicles, some thin discs, imaged end-on, could be seen, particularly in the CPZ liposome sample (Fig. 1B). Such disc-like structures have already been observed in liposomes without choles- terol, containing DSPE-PEG$_{2000}$ and lysophospholipid (Ickenstein et al., 2003). In this case, the lysophospholipid was seen as a micelle-forming element that allowed the formation of discs at grain boundaries. In the present study, no lysolipid was present but it is possible that CPZ can interact with phospholipids, and this could be the driving force for disc formation. By forming complexes with CPZ, CDs reduced the formation of discs, particularly in the SBE-β-CD/CPZ and SGM/CPZ systems where there is a high affinity between the CD and the drug (Table 1) (Wang et al., 2020). CPZ would then not be available to interact with the liposome bilayer and many of the objects seen in the cryo-EM images for these CDs were indeed vesicles. However, in the case of the HP-γ-CD/ CPZ system, both discs and vesicles were observed, confirming the poor complexation capacity of this CD with respect to CPZ. It should be noted that in the case of SBE-β-CD/CPZ system, the vesicles obtained are very sensitive to the electron beam despite the low electron dose used (10 electrons per Å$^2$ per s).

An effect of HP-β-CD on the size of liposome preparations has already been reported by Maestrelli et al. for multilamellar liposomes encapsu- lating CD/ketoprofen complexes, which were larger than liposomes with the drug alone (Maestrelli et al., 2005; Maestrelli et al., 2006). Cav- alcanti et al. made similar observations with β-lapachone/HP-β-CD li- posomes (Cavalcanti et al., 2011). However, Piel et al. did not see any differences with betamethasone alone and with various CDs, including HP-γ-CD (Piel et al., 2006).

The zeta potential of the preparations was measured as an indication of the charge on the particle surface. Empty liposomes showed a value of -30 mV (Table 2). This could be the result of the anionic nature of the headgroup of DSPE-PEG$_{2000}$ which confers a net negative charge at pH 7.4 despite the PEG at the surface. However, liposomes containing CPZ without CD showed a less negative zeta potential of -16 mV. This may

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Cryo-electron microscopy of ready liposomes. Images of final dehydration-rehydration vesicle (DRV) formulations were obtained by cryo-EM (A-E). Zoom on DRV + CPZ (B) shows undulating bilayer, while zoom on DRV + HP-γ-CD/CPZ shows example of lipid discs. Scale bars represent 200 nm (A, D, E) or 400 nm (B, C).</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** HL-60 cells are unaffected by CDs and empty liposomes but sustain a dose-dependent reduction in viability when treated with CPZ and CD/CPZ liposome formulations. HL-60 cells were measured for viability relative to control after incubation for 48 h with either: A) cyclodextrins (CD), B) empty dehydration-rehydration vesicles (DRV), or C) either free chlorpromazine (CPZ), CPZ-loaded DRV, or DRV loaded with CD/CPZ complexes. The results are shown as the mean ± SD of four wells measured by metabolic conversion of the MTS reagent.</figcaption></figure>

indicate some CPZ presence in the liposome membrane. As far as the DCL formulations were concerned, the values were intermediate, ranging from -19 to -28 mV. It was notable that there was an approximate correlation between the EE and the zeta potential, suggesting that even when CPZ is associated with CD, some interaction with the cell membrane could occur. It should be noted, however, that zeta potential measurements were made in low conducting medium and the charge at the liposome surface would be shielded in physiological media with a higher ionic strength.

The photodegradation of CPZ, both free and encapsulated in liposomes, was investigated revealing nearly complete degradation after 240 min, with an initial enhanced degradation rate in liposomes, but a reduced rate over time, resulting in 1.4 ± 0.7% residual CPZ (see Supplementary Materials).

## 3.2. Cytotoxicity of free and encapsulated CPZ

HL-60 promyelocytic acute myeloid leukaemia cells were used to assess the effect of unloaded and non-encapsulated CDs and unloaded/empty liposomes after a 48-hour incubation. No decrease in cell viability was observed after incubating with any of the three CDs at concentrations up to 100 µM (Fig. 2A), or with empty liposomes at lipid concentrations up to 1.6 mM (Fig. 2B). These findings are in accordance with many other observations of the safety of CDs for pharmaceutical applications (Loftsson et al., 2005), which generally present no cytotoxicity at the concentrations used in pharmaceutical preparations. For example, the IC₅₀ concentrations of HP-β-CD and ionic β-CDs were found to be higher than 200 mM against Caco-2 cells (Kiss et al., 2010). Although the methylated-β-CD derivatives are more toxic, most of them have an IC₅₀ higher than 50 mM. In fact, CD complexation can reduce the toxicity of a drug: chlorzoxazone in complex form with β-CD and HP-β-CD proved less toxic to normal liver cells compared to the free form (Tang et al., 2015). As far as in-vivo toxicity is concerned, the LD₅₀ doses of intravenously administered CD are quite high: 1.0 g per kg, 0.79 g per kg, and 4.0 g per kg for α-, β-, and γ-CD, respectively (Davis & Brewster, 2004).

Previous findings demonstrated that free, i.e. unencapsulated, CPZ exhibits cytotoxicity towards both AML cell lines and primary blasts from AML patients (Gundersen et al., 2022). To determine whether encapsulated CPZ retained the same potency as the free drug, all formulations were tested against four different AML cell lines with distinct cytogenetic profiles commonly observed in AML. HL-60, which is deficient for the tumour suppressor p53, MOLM-13 which is heterozygous for the poor prognostic mutation FLT-3 internal tandem duplication (ITD), and MV4-11, which is homozygous for FLT-3-ITD, and finally OCI-AML3, which has multiple mutations and complex karyotype. In line with our previous findings (Gundersen et al., 2022), free CPZ had an EC₅₀ of between 12 and 14 µM after 24 h of incubation with AML cells (Fig. 3). With increasing incubation time up to 72 h, the EC₅₀ values decreased to between 7 and 10 µM (Fig. 3). For HL-60 cells after 48 h,

EC₅₀ of free CPZ was 11.9 µM (Fig. 2C). The effect towards normal rat kidney epithelial cell line NRK was noticeably lower, with EC₅₀ of 49 µM at 24 h, which decreased to 22 µM at 72 h of incubation.

The affinity of CPZ towards the different CDs used has been shown to vary (Table 1) (Wang et al., 2020). These differences could also influence their ability to deliver CPZ as a free form into the cells, but also to avoid off-target effects on normal cells and tissues. The cytotoxic potential of the different liposomal formulations varied, with CPZ- and HP-γ-CD/CPZ-loaded liposomes demonstrating both similar and the highest cytotoxicity among all formulations. In contrast, SBE-β-CD/CPZ-loaded liposomes were less potent, and SGM/CPZ-loaded liposomes showed no reduction in viability after 24 or 48 h, even at CPZ concentrations up to 100 µM (Fig. 2C and 3).

Increasing the incubation time to 72 h resulted in lower EC₅₀ values for the formulations, demonstrating a slight time-dependent cytotoxicity (Fig. 3). Importantly, even after 72 h, empty liposomes without CD did not affect the viability of the cells (Fig. 3), nor did liposomes with only CD (data not shown).

These results suggest that CPZ complexed with CD is released more slowly from the liposomes and is therefore less available to the AML cells. Notably, the activity towards the cells is inversely proportional to the affinity of the CDs to CPZ determined by isothermal titration calorimetry in previous work: SGM > SBE-β-CD > HP-γ-CD (Wang et al., 2020). In fact, to our knowledge the SGM/CPZ complex has one of the highest binding constants that has ever been reported of a CD, at 6.37 × 10⁹ M⁻². These results provide evidence for a delayed release mechanism for the DCL systems. Our attempt to directly demonstrate different chlorpromazine release profiles as a function of the various DCLs tested was unsuccessful due to the retention of chlorpromazine on the materials used in the experiments (dialysis tubes and ultrafiltration devices).

The effect of CPZ on the normal rat kidney (NRK) cells was less marked than that of the different AML cell lines for both 24- and 72-hour incubations (Fig. 3). However, unlike for AML cells, liposomal formulations did not demonstrate a reduced availability of CPZ on the normal NRK control cells. After 24 h of incubation, liposomes loaded with either CPZ or HP-γ-CD/CPZ appeared more toxic towards the NRK cells compared with free CPZ, with EC₅₀ values of 30 and 35 µM, respectively, compared to 49 µM of free CPZ (Fig. 3). The inverse relationship between cytotoxic activity and affinity constant of the CD/CPZ pairings shown for AML cells was also observed for NRK, with SBE-β-CD/CPZ liposomes showing a lower cytotoxicity with an EC₅₀ of 51 µM, and SGM/CPZ liposomes showing no cytotoxicity for any of the tested concentrations (Fig. 3). Empty liposomes proved equally non-toxic. However, a stronger time-dependent toxicity of liposomal CPZ was observed in NRK cells than in AML cells. After 72 h of incubation, the NRK toxicity of all the DCL formulations were much more marked, and furthermore, the EC₅₀ values of all the formulations were lower than that of CPZ alone at 72 h (Fig. 3). Despite higher EC₅₀ values at 24 h compared to AML cell lines, NRK incubated for 72 h were either equally or more sensitive to liposomal formulations than AML cells (Fig. 3). It has been shown that
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** DRV formulations demonstrate dose-dependent cytotoxicity towards AML and normal cell lines after various incubation times. The AML cell lines MOLM-13, OCI-AML3, and MV4-11, and the NRK cell line, were measured for viability after incubation with various concentrations of either empty dehydration-rehydration vesicles (DRV), free chlorpromazine (CPZ), CPZ-loaded DRV, or DRV loaded with different cyclodextrin (CD)/CPZ-complexes. Cells were either incubated for 24 or 72 h. The results are shown as viability relative to untreated cells as measured by the metabolic conversion of the WST-1 reagent and represent the mean ± SD from 2 to 5 wells obtained from two separate experiments.</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>**Fig. 4.** DRV formulations demonstrate CD-dependent cell death kinetics. MOLM-13 cells were incubated for up to 12 h with 100 µM chlorpromazine (CPZ) in either free or encapsulated form, or empty (dehydration-rehydration vesicles) DRV of equivalent lipid concentration, before being fixed and stained with 2 % formaldehyde (in pH 7.4 PBS) containing 0.005 mg/mL Hoechst33342. Nuclei-stained cells were imaged with an Axiovert 200M fluorescence microscopy and counted for apoptosis. The results are shown as the mean ± SD of a triplicate experiment.</figcaption></figure>

dopamine and serotonin inhibit apoptosis in normal cells, including NRK (Talaei et al., 2011), and it can be that our DCL formulations enhanced an anti-dopamine, pro-apoptopic effect on these cells, either by protecting CPZ from degradation, or by increasing cellular uptake. Our results suggest that the NRK cells either take up the liposomes and/or CPZ more slowly than leukaemia cells, or that they are less sensitive to unencapsulated CPZ, and that there could be some selectivity of the formulations or a liposome-mediated increase in CPZ sensitivity for NRK cells. While the cause for this time-dependence against NRK is unclear, that our DCL formulations show some toxicity to the normal NRK cells highlights the need for surface-bound ligands, such as folate or monoclonal antibodies, that increases the selectivity against AML cells and reduces off-target effects during long-term exposure (Singh et al., 2019; Villaverde & Baeza, 2019).

To shed more light on any potential delayed release mechanism and time-dependent cytotoxicity of different DCL formulations, MOLM-13 cells were incubated with the different formulations adjusted to 100 µM CPZ, and apoptosis determined at different incubation times up to 12 h (Fig. 4). DRV formulations of both CPZ and HP-γ-CD/CPZ were equally available to MOLM-13 as free CPZ, as all three treatments resulted in 100 % cell death after 6 h with similar change in death over time. SBE-β-CD/CPZ liposomes however were slower to induce cell death, requiring 12 h to reach 100 % cell death, whereas SGM/CPZ liposomes after 12 h only resulted in around 20 % cell death. Empty liposomes of equivalent lipid concentration to the most concentrated DRV formulation (DRV + SGM/CPZ), did not cause significant cell death (Figs. 2 and 3), even at a 20 % v/v concentration. These results explain the results of earlier cytotoxicity experiments wherein the EC₅₀ varied based on incubation time and the CD included in the DCL formulation, which suggested that the CD-complexation could reduce the intracellular release of CPZ from the liposomes and limit its availability to the cells. Again, the findings correlate to the different binding strength of the various CD/CPZ complexes (Table1) (Wang et al., 2020).

These delayed release properties would be an advantage for maintaining an effective concentration of CPZ for a longer time after intravenous administration, or to delay drug release until the liposomes had entered the bone marrow. A similar phenomenon was observed by Chen et al., who found that the hydrophobic drug indomethacin was released rapidly from the lipid bilayers, whereas for liposomes containing CD/indomethacin complexes, a slower drug release was observed (Chen et al., 2007).

According to the authors, two routes of drug release from the DCL system can be envisaged: 1) the CD/drug complexes can be released from the liposomes intact by passing through the lipid bilayers; 2) free drug is released from the interior aqueous phase of liposomes as equilibrium is established between CD and drug. Therefore, in both situations, the rate of drug release will be affected by the affinity between the drug and the CD. Piel et al. reported slower release of beta-methasone into Carbopol gels from DCL formulations than from simple liposomes at rate that was related to the ability of the CD to complex the drug (Piel et al., 2006). In our results reported above, the strong delaying effect of SGM, related to the very high binding constant of the complex, is very promising for the development of DCL systems that will retain CPZ long enough for it to exert anti-AML effect without affecting the CNS.

### 3.3. Uptake of DRV in AML cells

To study cellular uptake of liposomes, rhodamine-labelled fluorescent dehydration-rehydration vesicle (F-DRV) formulations with or without CDs, but always without CPZ, were incubated with MOLM-13 cells for up to 8 h. CPZ was not included as its presence in fluorescent liposomes strongly quenched the fluorescence signal, resulting in unreliable data (not shown). The presence of free CDs did not affect the fluorescence of F-DRV, neither in terms of excitation spectra nor emission spectra (Supplementary Fig. S3). Cells incubated for 2 h were imaged with confocal microscopy to localise the liposomes, while cells incubated for various time points were measured for fluorescence intensity by flow cytometry. This approach enabled the determination of whether differences in cellular uptake could account for variations in cytotoxicity and kinetics among the formulations.

The flow cytometric data is presented in two graphs (Fig. 5A and B) that both show fluorescence intensities as given by the instrument after having been adjusted for the autofluorescence of control and the fluorescence of samples incubated for 1 min. The formulations differed in their fluorescence intensity, thus Fig. 5B shows data additionally adjusted for the differences in fluorescence of each formulation.

From the measured fluorescence intensities in Fig. 5A, the formulations share a similar uptake capacity, initial uptake rate, and time required for uptake capacity of around 2 h. SBE-β-CD-loaded liposomes differ slightly with a slower initial uptake rate and requires closer to 4 h to reach uptake capacity. However, while the volume of added liposomes was kept the same between formulations, two assumptions are made for this interpretation. The first assumption is an equal lipid concentration between formulations, and therefore equal concentration of the PE-conjugated rhodamine probe. This was not accounted for given the similar lipid concentrations between formulations, ranging from 23.1 to 28.4 mM. The second assumption was an equal fluorescence output from each formulation, but this varied greatly between formulations, as seen by spectrophotometric measurements in PBS buffer with 5 % Tween® 20 detergent (Supplementary Fig. S4). This difference in fluorescence output was therefore used to adjust the flow cytometry measurements of cellular fluorescence intensities in Fig. 3B.

Considering the adjusted fluorescence intensity values, the formulations share the same initial uptake rate and, apart from SBE-β-CD-loaded liposomes, also share an identical uptake capacity in MOLM-13 cells. SBE-β-CD-loaded liposomes again stand out, this time with a higher uptake capacity. One explanation could be that SBE-β-CD increases uptake by binding particularly well to PEG, thus reducing the presence of PEG on surface and therefore its “stealth” effect. Our findings demonstrate that, if not accounted for, variations between formulations can skew measurements and misrepresent the uptake rate and saturation.

As measured by flow cytometry (Fig. 5A, B) and visualised by confocal microscopy (Fig. 5C-G), the fluorescent DRV quickly undergo cellular uptake into MOLM-13 cells. Uptake saturation of the
<figure><img src="image_8.png" /><figcaption>**Fig. 5.** DRV formulations are quickly internalised into MOLM-13 AML cells. MOLM-13 cells were incubated with rhodamine-labelled fluorescent dehydration-rehydration vesicles (F-DRV), that were either empty or loaded with cyclodextrins (CD), at 2 % v/v concentration for up to 8 h. After incubation, the cells were washed twice in PBS and their red fluorescent intensity analysed using an Accuri C6 flow cytometer (A, B). At least 10,000 events were collected, and the median fluorescence from each sample was determined. See Supplementary Fig. S2 for examples of the gating strategy. The results represent the mean ± SD of either two experiments (F-DRV, F-DRV + SGM) or a single experiment (F-DRV + HP-γ-CD, F-DRV + SBE-β-CD), all performed in triplicates. Results are provided as direct measurements from the flow cytometer (A) or adjusted for the relative fluorescence intensity of each formulation (B). Cells incubated for 2 h were also washed twice in PBS, fixed in 2 % buffered formaldehyde (pH 7.4 PBS), and imaged with an Andor Dragonfly 505 confocal microscope fitted with a CFI SR HP Apo TIRF 100× objective (C-G). Scale bars in the confocal micrographs represent 15 µm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

formulations was reached within 2–4 h of incubation, and there were only minor differences in uptake between the different formulations, suggesting that the CDs do not significantly influence liposome uptake. Therefore, differences in internalisation cannot account for the differences in cytotoxicity between formulations with different CDs.

Given the similar uptake behaviour of the formulations and how quickly uptake saturation is reached for MOLM-13, any difference in cytotoxicity between 24 and 72 h is unlikely to be accounted for by a difference in the rate of liposome uptake. A more likely reason for the difference in cytotoxicity is CPZ being released intracellularly at different rates based on the CD and its affinity for CPZ. The observed uptake saturation also does not suggest the complete uptake of all the liposomes present in the medium, but a combination of restricted uptake and surface-bound liposomes. In any case, these results demonstrate that the liposomes are internalised by MOLM-13, and further support the theory that differences in cytotoxicity between the formulations is not due to differences in liposomal uptake, but CD/CPZ affinity and subsequently the availability of free CPZ.

### 3.4. Biodistribution in zebrafish larvae and biocompatibility of liposomes

After being injected with fluorescent liposomes at two days post-fertilisation (dpf), zebrafish larvae were imaged for fluorescence by confocal microscopy within three hours post-injection (hpi), i.e. zero

days post-injection (dpi), and then at one- and two-days post-injection (Fig. 6). The injected F-DRV were either empty (Fig. 6C) or loaded with one of the three CDs (Fig. 6D-F), but no significant differences were observed between the formulations in their biodistribution. All formulations were observed circulating through the blood vessels, not only immediately after injection, but also at two days post-injection.

The 3D representations of the blood vessels in the brain at one day post-injection (Fig. 6G-J) show that liposomes are found in the blood vessels of the head and eyes, but their absence in the brain demonstrates that the liposomes do not cross the blood-brain barrier (BBB) of zebrafish larvae by endocytosis or passive transport. Indeed, it has been shown that the BBB of zebrafish shares structural and functional similarities to that of mammals, and that it matures in larval zebrafish between three and ten days post-fertilisation (Fleming et al., 2013). Already as early as 2.5–3 days post-fertilisation, the BBB shows reduced permeability and partial functionality for larger molecular weight molecules of 4 kDa and above (Fleming et al., 2013; Quiñonez-Silvero et al., 2020). With zebrafish models already established for a range of BBB- and CNS-related diseases, it is clear they can provide insights into BBB functions that have relevance for humans, and therefore utility in pre-clinical drug development (Quiñonez-Silvero et al., 2020).

In contrast to the brain, a substantial fluorescence signal was detected in the yolk sac of the larvae injected with various liposomes, with no fluorescence present in untreated control larvae. This indicates
<figure><img src="image_9.png" /><figcaption>**Fig. 6.** Distribution of DRV formulations after intravenous injection into zebrafish larvae. Zebrafish larvae were injected into the posterior cardinal vein at two days post-fertilisation (dpf) with approximately 4 nL of different rhodamine-labelled, fluorescent dehydration-rehydration vesicles (F-DRV). Zebrafish larvae were kept in fresh E3 medium at 28.5 °C and anesthetised in a 0.7 mM tricaine solution before injection and during imaging. Images were acquired at the day of injection, or zero days post-injection (dpi), and at one and two dpi using an Andor Dragonfly 505 confocal microscope fitted with a CFI Plan Apochromat Lambda 10× objective, 561 nm excitation laser, and 600/25 nm band pass filter. Fluorescence images of whole larvae (A-F) are a Z-projection obtained by summation of overlying pixels in the confocal Z-stack. Images also show large sections of the head of one dpi zebrafish larvae, with 3D representations of the blood vessels in the brain (G-J) obtained by pre-processing using a 3D-median filter and rendered using the ImageJ 3D Viewer plugin. All image processing was performed in ImageJ ver. 1.52p. The scale bars represent 500 µm (A-F) and 250 µm (G-J).</figcaption></figure>

that either F-DRV or the fluorescently labelled lipids are gradually accumulated in the yolk sac. It should be noted that the fluorescence images are made by summing the overlying pixels in the confocal Z-stack, and that this contributes to high signal within the large yolk sac compared to the miniscule blood vessels.

Another site of F-DRV accumulation is along the ventral caudal region of the larvae, which corresponds to the caudal haematopoietic tissue, in which macrophages and neutrophils undergo maturation. An explanation for this accumulation could be the discontinued endothelium of blood vessels from insufficient maturation. This area is crowded with maturing leukocytes of high phagocytic capacity (Rosowski, 2020). However, despite this accumulation of liposomes, as well as the accumulation in the yolk sac, liposomes were observed circulating in the blood vessels for at least 48 h after injection.

Zebrafish larvae exposed in water to various concentrations of free CPZ, CPZ-loaded DRV with or without CDs showed dose-dependent toxicity, with CPZ- and HP-γ-CD/CPZ-loaded DRV being most toxic, while SBE-β-CD and SGM/CPZ-loaded DRV provided better protection, highlighting the potential of zebrafish for toxicology research on drugs and nanocarriers (see Supplementary Materials).

The deposition of complement proteins on the surface of liposomes

after activation by the alternative pathway is the first step towards their recognition and phagocytosis by macrophages (Moghimi et al., 2019). Thus, for the DCL system to circulate in the blood, they should be poor activators of complement. This property was assessed by a miniaturised two-dimensional immunoelectrophoresis method that monitors the cleavage of the complement protein C3 (Coty et al., 2017). The results shown in Table 3 demonstrate that the electrophoretic profiles from serum incubated with the different DCL formulations were not significantly different to the spontaneous activation shown by serum incubated with HEPES buffer or Veronal-buffered saline (VBS²⁺), with only small peaks corresponding to the cleavage product and opsonin C3b. On the other hand, serum incubated with a known activator such as poly(isobutylcyanoacrylate) nanoparticles (Coty et al., 2017), here denoted as A1 nanoparticles, showed a high degree of conversion. Typical electrophoretic profiles are shown in Supplementary Fig. S6. These results allow us to predict that the DCL systems could persist in the circulation and reach the bone marrow, and they are in accordance with our observations in zebrafish larvae, since if complement activation had occurred, the liposomes would have been rapidly cleared from the circulation. The role of PEGylation of liposomes has been demonstrated in earlier studies in zebrafish, where PEGylated liposomes were found in the circulation 72 h after injection, whereas non-PEGylated liposomes were absent from circulation less than 24 h after injection (Evensen et al., 2016). These results highlight the potential of repurposing CPZ for anti-leukaemic therapy using liposomal drug carriers.

<caption>**Table 3**</caption>

Drug-in-cyclodextrin-in-liposome formulations show minimal effect on complement activation.

<table><thead><tr><th>Sample</th><th>C3 cleavage (%)</th></tr></thead><tbody><tr><td>VBS<sup>2+</sup></td><td>15.5 ± 2.6</td></tr><tr><td>HEPES buffer</td><td>14.3 ± 0.9</td></tr><tr><td>Empty DRV</td><td>11.2 ± 4.6</td></tr><tr><td>CPZ-loaded DRV</td><td>15.9 ± 2.6</td></tr><tr><td>HP-γ-CD/CPZ-loaded DRV</td><td>11.2 ± 2.1</td></tr><tr><td>SBE-β-CD/CPZ-loaded DRV</td><td>11.7 ± 3.3</td></tr><tr><td>SGM/CPZ-loaded DRV</td><td>11.4 ± 2.0</td></tr><tr><td>A1 nanoparticles (positive control)</td><td>87.4 ± 0.4</td></tr></tbody></table>

## 4. Conclusion

The results reported above demonstrate that it is possible to encapsulate CD/CPZ complexes within liposomes that are suitable for intravenous administration because of their small size and low complement-activating properties. Due to the low toxicity of the liposome form, it was possible to evaluate the effect of encapsulated CPZ against a panel of
cell lines. Encapsulated in a drug-in-CD-in-liposome system, CPZ exhibited activity towards leukaemic cells, showing some selectivity over normal cells. The encapsulation of CPZ delayed the effect compared with free drug and showed CD-dependent cytotoxicity that correlates with CD/CPZ binding affinity. In MOLM-13 AML cells, liposomes were shown to be quickly internalised, and in zebrafish larvae, liposomes were observed to not cross the BBB and accumulate in the brain. Taken together, these results demonstrate the potential of the drug-in-CD-in-liposome delivery system to ensure delivery of CPZ to leukaemic cells while sparing the central nervous system during AML chemotherapy. The addition of a targeting agent to the surface of the liposomes should further improve the specificity of action (Myhren et al., 2014).

### CRediT authorship contribution statement

**Edvin Tang Gundersen:** Writing – review & editing, Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Zhiqiang Wang:** Writing – review & editing, Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation. **Jan-Lukas Førde:** Writing – original draft, Visualization, Methodology, Investigation. **Eric Larquet:** Writing – review & editing, Visualization, Investigation. **Reidun Æsøy:** Investigation, Data curation. **Hugo Roussel:** Visualization, Investigation, Formal analysis. **Lou Tosi:** Visualization, Investigation, Formal analysis. **Gillian Barratt:** Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization. **Lars Herfindal:** Writing – review & editing, Supervision, Resources, Project administration, Methodology, Funding acquisition, Conceptualization. **François-Xavier Legrand:** Writing – review & editing, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization.

### Funding sources

This work was funded by grants from the Western Norway Health Authorities to ETG (grant no.: 912122), LH (grant no.: 912052), JLF (grant no.: F-12533), from the Norwegian Cancer Society to LH (grant no.: 206696), and from the Norwegian Society for Children's Cancer to LH and RA (grant no.: 190004 and 230001). The funding body did not take part in study design, data interpretation, or writing of the manuscript. This work was also supported by the Laboratory of Excellence in Research on Medication and Innovative Therapeutics (LERMIT, grant number: ANR-10-LABX-33) and the China Scholarship Council with a Ph.D. grant to ZW.

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgements

We thank Pr. Luc de Chaisemartin (Hôpital Bichat-Claude Bernard, AP-HP) for providing Sugammadex and Dr. Christine Vauthier (Institut Galien Paris-Saclay) for her assistance in determining complement activation. Ashland Global Specialty Chemicals Inc. and Ligand Pharmaceuticals Inc. are gratefully acknowledged for generous gifts of HP-γ-CD and SBE-β-CD, respectively.

### Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.carbpol.2025.123478.

### Data availability

Data will be made available on request.

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