phenomenon to break up a liquid film into fine droplets. In this case, the phenomenology of the droplets formation process can be explained with cavitation‐wave hypothesis or with the capillary‐wave hypothesis [23] which analyse the behaviour of a liquid on the solid surface vibrating based on Taylor instability criteria [24]. When the vibration has a frequency of 30 Hz, above a threshold value of amplitude, on the liquid surface capillary waves are formed consisting in crests and troughs. At higher values of amplitude, liquid droplets, separated

In general, ultrasonication is an easy and scalable process which shows competitive energy costs, applicable in a vast range of fields and for disparate applications. Regarding the particle size reduction, it takes advantage from ultrasound process of the ability to break up lipid structures. In case of nanoliposomes production, the energy is used to break the lipid bilayer into smaller pieces; then these pieces close themselves in spherical structures producing SUVs. In comparison with other sizing techniques, the ultrasonic, which once shows great potential due to reduced time spent and easiness in use, is a simple and efficient method to reduce liposome dimension [26, 27]. Ultrasonic process does not require a number of passages of liposome suspension through a membrane such as extrusion method, and high pressures are not required. Furthermore, the final dimensions of particles are not fixed to the pore size of the membrane, but it is possible to change the vesicle size according to the application requirements. This is possible by controlling the duty cycle, the discontinuous process before described in which the liquid sample is sonicated in periodic time intervals followed by switch off in energy supply. Finally, due to the critical importance to have a sterile environment during loaded SUVs production, another advantage is that sonotrode tip is simpler to

As described in Ref. [28], from a thermodynamics point of view, the free energy of the liposome membrane is mainly given by two contributions: the elastic energy due to membrane curvature and the tension energy due to the edge of the layer. Indeed, at the boundary of a bilayer, the polar heads have to be arranged as a semi‐circle, in order to connect the two monolayers [29]. The elastic energy has been estimated by Helfrich [30] as proportional to the second power of twice the mean surface curvature, 1/*R* [m−1], and to Gaussian surface curva-

to be proportional to the length of the bilayer edge [30, 31], by the edge tension parameter, *γ* [J m−1]. On the basis of simple geometrical reasoning [31], a disc initially of radius *ρ*D [m], with

Therefore, the thermodynamic of the process could be described once the number and the size of the starting discs, *N* and *ρ*D, as well as the curvature of the vesicle, *1/R*, are known;

[m] of *R*S

\_1 2(*kc* + \_1 2 *k*¯ *c*) 4*π ρD* 2 \_ *R*2 ] + [

[J] and *k*¯¯*c*

). Summarizing, for *N* vesicles with a mean curvature *1/R*, the total free energy

, which is (partially) bended towards forming a sphere (equivalent in

 *= ρ*D */2*), has an edge length equal to *L* = (2π*ρ*D

*γ*(2*π ρD*)√

[J], the edge energy are assumed

\_

1 − *ρD* 2 \_

4 *R*2]} (1)

2 )

from the wave crest, are formed [25].

34 Liposomes

clean and sterilize [4].

a surface area *A*D = π*ρ*D

area, then with a sphere radius *R*S

could be estimated by Eq. (1) [1]

*g* = *N*(*g*elastic + *g*edge) = *N*{[

ture, 1/*R*2

√1−(*ρ*D 2 /(4*R*2

*2.1.3. Phenomenological aspects*

[m−2]; through the main elastic moduli, *k*c

2

**Figure 3** shows a schematization of the developed simil‐microfluidic apparatus. The setup is constituted by five main process sections: a feeding section, a pumping section, a production section, a homogenization section and a recovery section.

**Figure 3.** Nanoliposome production through the simil‐microfluidic setup. The main sections are reported: feeding, pumping, production, homogenization and recovery. From Tanks [1–2], lipids/ethanol (eventually containing the hydrophobic molecule to be encapsulated) and water (eventually containing the hydrophilic molecule to be encapsulated) solutions are pushed, through peristaltic pumps (Pumps 1–2), to the production section. Here, after the lipid solution injection into the polar phase, nanometric vesicles are formed. The hydroalcoholic solution is then recovered and homogenized through an ultrasound‐assisted process. Finally, the suspension is recovered and characterized.

In particular, the feeding section consists in two lines. The first one is made up by a stirred tank filled with lipids/ethanol solution (Tank 1 of **Figure 3**), in which a hydrophobic active molecule to be encapsulated, conveyed in a silicon tube with an internal diameter of about 2 mm, is eventually dissolved. The second line includes a stirred tank filled with the hydration solution (Tank 2 of **Figure 3**), which can be pure water or an aqueous solution, eventually containing the hydrophilic active molecule to be encapsulated, conveyed in a flexible silicone tube with an internal diameter of few millimetres.

The pumping section is composed by two single‐head peristaltic pumps (Verderflex OEM mod. Au EZ) indicated as Pumps 1–2 in **Figure 3**.

The lipids/ethanol solution tube ends with several tenths of millimetres internal diameter needle inserted into a silicon tube, an extension of the water tube. This is the production section sketched in **Figure 3**. In this section, an interdiffusion of the two pushed liquids occurs leading to the formation of liposomes on nanometric scale; the suspension is then collected inside a tank and subjected to a homogenization in order to optimize vesicles size distribution. The suspension is finally recovered and characterized.

Briefly, the process starts when lipids/ethanol and water solutions are pushed through peristaltic pumps into the production section, where liposomal vesicles are formed directly on nanometric size (the phenomenology behind vesicles formation through a simil‐microfluidic approach will be discussed in the next paragraph). The formed hydroalcoholic solution is recovered and subjected to a homogenization process through the duty cycle sonication protocol, previously described for the ultrasound‐assisted thin‐film hydration method [28] (Section 2.1.1).

#### *2.2.2. Phenomenological aspects*

In particular, the feeding section consists in two lines. The first one is made up by a stirred tank filled with lipids/ethanol solution (Tank 1 of **Figure 3**), in which a hydrophobic active molecule to be encapsulated, conveyed in a silicon tube with an internal diameter of about 2 mm, is eventually dissolved. The second line includes a stirred tank filled with the hydration solution (Tank 2 of **Figure 3**), which can be pure water or an aqueous solution, eventually containing the hydrophilic active molecule to be encapsulated, conveyed in a flexible silicone

**Figure 3.** Nanoliposome production through the simil‐microfluidic setup. The main sections are reported: feeding, pumping, production, homogenization and recovery. From Tanks [1–2], lipids/ethanol (eventually containing the hydrophobic molecule to be encapsulated) and water (eventually containing the hydrophilic molecule to be encapsulated) solutions are pushed, through peristaltic pumps (Pumps 1–2), to the production section. Here, after the lipid solution injection into the polar phase, nanometric vesicles are formed. The hydroalcoholic solution is then recovered and homogenized through an ultrasound‐assisted process. Finally, the suspension is recovered and characterized.

The pumping section is composed by two single‐head peristaltic pumps (Verderflex OEM

The lipids/ethanol solution tube ends with several tenths of millimetres internal diameter needle inserted into a silicon tube, an extension of the water tube. This is the production section sketched in **Figure 3**. In this section, an interdiffusion of the two pushed liquids occurs leading to the formation of liposomes on nanometric scale; the suspension is then collected inside a tank and subjected to a homogenization in order to optimize vesicles size distribu-

tube with an internal diameter of few millimetres.

36 Liposomes

mod. Au EZ) indicated as Pumps 1–2 in **Figure 3**.

tion. The suspension is finally recovered and characterized.

From a phenomenological point of view, liposome formation is governed by the molecular diffusion between two phases: the organic solvent, in which the lipids are solubilized, and the water; the latter simultaneously diffuses into the organic solvent in order to reduce its concentration below the critical value required for the lipid's solubilization. During the diffusion process, lipid vesicles on nanometric scale start to form through a mechanism called 'self‐assembly', according to the theory by Lasic and Papahadjopoulos [32]: lipids dissolved in an organic solvent are in the form of bilayer fragments (phospholipid bilayer fragments, BPFs), the interdiffusion of the water and the organic solvent reduces the solubility of the lipids in the solvent causing thermodynamic instability of BPF edges, inducing the curvature and the closure of bilayer fragments which allow the formation of liposomal vesicles [32]. In the simil‐microfluidic setup developed, through the use of constructive expedients (millimetric tubes, peristaltic pumps and injection needle), the reproduction of the laminar flow regime was possible, all the Hagen‐Poiseuille assumptions being satisfied, that is, the Reynolds number was found to be less than 2100 for all the volumetric flow rate conditions tested; the piping length in which the two phases interdiffuse was longer than the 'entrance length' required to obtain the parabolic profile [33]. In particular, for a microfluidic system and thus for a laminar flow, liposome formation occurs at the interfaces between the alcoholic and water phases, when they start to interdiffuse in a direction normal to the liquid flow stream. Changes in flow conditions result in size variations of the insertion section of the organic phase reflecting on the vesicles dimensional features. In particular, increasing the volumetric flow rates ratio, the size of the insertion section of the organic phase decreases; this leads to a major dilution of the organic phase limiting the formation of long BPFs, thus inducing the production of liposomal structures of small dimensions. In general, it was shown that the variation in shear forces at the interface of the two fluids has no consequence on liposome structure. In particular, maintaining constant the volumetric flow rates ratio and changing both the buffer and the lipid alcoholic solutions volumetric flow rates, Jahn and collaborators have demonstrated that it is not the magnitude of the shear forces between the parallel‐layered stream in having significant impact on liposome's size and size distribution but the stream width (which depends on the volumetric flow rates ratio) [20]. Due to the developed apparatus, the phenomenology connected to the vesicles formation through a microfluidic approach was achieved, exceeding the limit of the bulk methods where a driving force of entropic nature leads to the liposome formation. Local fluctuations of the lipid concentration in a bulk solution make difficult to control the size and the polydispersity of the produced vesicles. On the contrary, the presence of intubated laminar flow with the relative matter diffusive transport allows to minimize the fluctuations of the lipid concentration inside the tubes and to modulate the size and the size distribution of the final vesicles.

#### *2.2.3. Influence of process parameters on liposome dimensional features*

When using the simil‐microfluidic approach, the lipid concentration and the volumetric flow rate ratio [Vhs/Vls] have a great influence on liposome's size distribution as described in Ref. [34] and also found by Jahn and collaborators for a microfluidic hydrodynamic‐focusing platform [35]. In particular, increasing the ratio between the water volumetric flow rate to the lipids‐ethanol volumetric flow rate, the PDI value increases as shown in **Figure 4B**. On the contrary, the effect of the ultrasound‐assisted process in reducing PDI and thus in ameliorating their size distribution (homogenizing) can be observed (**Figure 4B**).

Another crucial parameter affecting nanoliposome's dimensional features is the lipid concentration. In particular, as visible in **Figure 5A**, increasing lipid concentration, the liposome diameter also increases. This can be explained by the fact that at equal fluid dynamic conditions, a greater number of lipids impact at the same alcohol/water interface area dissolving in the same water volume, thus joining to form larger vesicles. The sample seems to be better homogenized at the higher‐tested lipid concentration (**Figure 5B**).

**Figure 4.** Liposome diameter size (A) and polydispersity index (PDI) (B) before and after sonication treatments at different volumetric flow rates [34].

**Figure 5.** Liposome diameter size (A) and polydispersity index (PDI) (B) before and after sonication treatments at different phosphatidylcholine (PC) concentrations in the hydroalcoholic solution [34].

## **3. Case histories**

*2.2.3. Influence of process parameters on liposome dimensional features*

ing their size distribution (homogenizing) can be observed (**Figure 4B**).

homogenized at the higher‐tested lipid concentration (**Figure 5B**).

different volumetric flow rates [34].

38 Liposomes

When using the simil‐microfluidic approach, the lipid concentration and the volumetric flow rate ratio [Vhs/Vls] have a great influence on liposome's size distribution as described in Ref. [34] and also found by Jahn and collaborators for a microfluidic hydrodynamic‐focusing platform [35]. In particular, increasing the ratio between the water volumetric flow rate to the lipids‐ethanol volumetric flow rate, the PDI value increases as shown in **Figure 4B**. On the contrary, the effect of the ultrasound‐assisted process in reducing PDI and thus in ameliorat-

Another crucial parameter affecting nanoliposome's dimensional features is the lipid concentration. In particular, as visible in **Figure 5A**, increasing lipid concentration, the liposome diameter also increases. This can be explained by the fact that at equal fluid dynamic conditions, a greater number of lipids impact at the same alcohol/water interface area dissolving in the same water volume, thus joining to form larger vesicles. The sample seems to be better

**Figure 5.** Liposome diameter size (A) and polydispersity index (PDI) (B) before and after sonication treatments at

**Figure 4.** Liposome diameter size (A) and polydispersity index (PDI) (B) before and after sonication treatments at

different phosphatidylcholine (PC) concentrations in the hydroalcoholic solution [34].

#### **3.1. Pharmaceutical application: nanoliposome vectors for siRNAs delivery**

The short‐interfering RNAs are double‐stranded RNA molecules able to target disease components, at genetic level that are considered 'undruggable' with the conventional medicines; thus, their use in the development of innovative gene therapies is growing faster in recent years. Due to their low stability in physiological fluids, low‐membrane permeability and their short half‐life in the circulatory system, siRNAs are not useable in their naked form and require to be encapsulated in suitable carriers.

As described in Ref. [36], siRNA sequences directed against E2F1 transcription factor (siE2F1‐1324) were encapsulated inside positively charged vesicles purposely designed and produced to enhance the interaction with both the negatively charged siRNA, improving its encapsulation efficiency, and the cell‐plasmatic membrane, also negatively charged, improving siRNA incorporation in the target cells [37, 38]. It was demonstrated that E2F1 promotes the aggressiveness of human colorectal cancer by activating the ribonucleotide reductase small subunit M2 whose high expression induces cancer and contributes to tumour growth and invasion [39]. Due to the observed correlation, the inhibition of E2F1 expression was studied as a potential way to treat colorectal cancer by encapsulating siE2F1 in cationic nanoliposomes suitably produced by the ultrasound‐assisted technique developed. The loaded nanoliposomes were then transfected in human cell lines and in intestinal human biopsy fragments (collected from IBD donors during lower endoscopy performed for colonic cancer screening) to investigate their in vitro and ex vivo silencing activity. In particular, siE2F1 nanoliposomes were transfected in HT29 human colon adenocarcinoma cell line, where conditions are more reproducible, and in cultured human biopsies, in which the cell‐cell interactions, thus the human intestinal mucosa cytoarchitecture, are preserved unlike isolated cell cultures.

#### *3.1.1. Materials and methods*

*Formulation:* Nanoliposomes loaded with siRNA sequences for E2F1 expression inhibition were designed and produced by using cholesterol (CAS 57‐88‐5), l‐α‐phosphatidylcholine from egg yolk (CAS 8002‐43‐5) and dioleoyloxy propyl‐N,N,N‐trimethylammonium propane (DOTAP) (CAS 132172‐63‐1, >99 % pure), purchased from Sigma‐Aldrich (Milan, Italy). In designing the liposome bilayer, the cationic DOTAP phospholipid was chosen to electrostatically interact with the negative siRNA molecules, promoting siRNA encapsulation inside the lipid vesicles, and to interact with the negative cell membrane. The charge ratio between DOTAP and siRNA sequences used was 8.5:1 (±), selected on the basis of previous work [40].

The siRNA sequence direct against E2F1, the siE2F1‐1324, was selected [41, 42]. siE2F1‐SUV complexes were produced using the thin‐film hydration method [16] followed by duty cycle sonication [28]. Briefly, PC, DOTAP and CHOL at 3:0.3:1 (mol:mol) ratio were dissolved in chloroform/methanol at 2:1 (vol/vol). The solvent was removed and the produced lipid film was hydrated with a phosphate buffer solution (PBS; potassium phosphate monobasic of 0.2 M, sodium hydroxide of 0.2 M, pH 7.4) containing siE2F1‐1324 at 8 μM. siRNA‐positive and ‐negative controls (scramble siRNA) were also encapsulated. Finally, the above‐described steps were followed also for unloaded liposome production with the only difference in the hydration solution which was pure PBS not containing siRNA sequences. By this way, multilamellar vesicles were achieved, maintained at room temperature for 2 h and then diluted obtaining an siRNA concentration of 4 μM for the loaded samples with a 1:260 (w/w) siRNA/total lipid ratio.

*Production:* In order to obtain nanoliposomes, samples were subjected to the duty cycle sonication process through the developed ultrasound‐based size reduction method and previously described (Section 2.1.1).

Unloaded and siRNA‐loaded SUVs were morphologically characterized through an optical microscope, equipped with software to capture the images in the fluorescence field (Axioplan 2‐ Image Zeiss, Jena, Germany). The size and zeta‐potential determinations were performed by using the ZetasizerNano ZS (Malvern, UK) with non‐invasive backscatter (NIBS) optics. The resulting particle size distribution was plotted as the number of liposomes versus size. The encapsulation efficiency (E.E.) by spectrophotometric (Lambda 25 UV/VIS Spectrophotometer, *λ* = 260 nm for RNA molecules) and electrophoretic assay (run on 1.5% agarose gel) was evaluated. The E.E. was determined as the percentage of siRNA encapsulated into SUVs, calculated subtracting the amount of siRNA present in the supernatant of the centrifuged sample from the total amount of siRNA included in the formulation, to the initial amount of siRNA used.

In order to evaluate siRNA‐nanoliposome cytotoxicity and their potential in E2F1 silencing, the complexes were transfected at 200 nM in human colorectal adenocarcinoma cell lines HT29 and in human colon mucosa biopsies, isolated from donors during colonoscopy.

#### *3.1.2. Results and discussion*

The production process (formulation and manufacturing) is of crucial importance in preparing siRNA‐liposome complexes with the desired shape and size without damaging siRNA's integrity. The shape of liposomes is the main factor affecting carrier's entry in the cellular compartment. Spherical nanoparticle's uptake in mammalian cells was demonstrated to be 3.75–5 times more than rod‐shape nanoparticles, indicating that the carrier's curvature can affect the entry in the plasma membrane [43]. In that regard, spherical SUVs were obtained through the ultrasound‐assisted method adopted which has been successfully used to produce stable siRNA‐SUV complexes on nanometric scale. The achieved siRNA‐liposomes, with a mean diameter size of 38 nm (**Table 1**), are useful for the EPR effect, which involves carrier's extravasation through tumour vascular fenestrations of 50–100‐nm range size. Another important feature is the surface charge of liposomes. Zeta‐potential (*ζ*) was investigated for both unloaded and SUVs encapsulating siRNA samples; the results are presented in **Table 1**. The positive zeta‐potential makes the produced liposomes applicable for the encapsulation of negative siRNA molecules and also promotes the fusion with the negatively charged cell membrane. The *ζ*‐value of the unloaded liposomes (27.90 ± 1.60) appears to be significantly higher than the zeta‐potential of the siE2F1‐1324‐SUV sample (18.02 ± 1.07 mV) suggesting that a strong complexation in addition to the siRNA core encapsulation occurred for the loaded structures.

sodium hydroxide of 0.2 M, pH 7.4) containing siE2F1‐1324 at 8 μM. siRNA‐positive and ‐negative controls (scramble siRNA) were also encapsulated. Finally, the above‐described steps were followed also for unloaded liposome production with the only difference in the hydration solution which was pure PBS not containing siRNA sequences. By this way, multilamellar vesicles were achieved, maintained at room temperature for 2 h and then diluted obtaining an siRNA concentration of 4 μM for the loaded samples with a 1:260

*Production:* In order to obtain nanoliposomes, samples were subjected to the duty cycle sonication process through the developed ultrasound‐based size reduction method and previously

Unloaded and siRNA‐loaded SUVs were morphologically characterized through an optical microscope, equipped with software to capture the images in the fluorescence field (Axioplan 2‐ Image Zeiss, Jena, Germany). The size and zeta‐potential determinations were performed by using the ZetasizerNano ZS (Malvern, UK) with non‐invasive backscatter (NIBS) optics. The resulting particle size distribution was plotted as the number of liposomes versus size. The encapsulation efficiency (E.E.) by spectrophotometric (Lambda 25 UV/VIS Spectrophotometer, *λ* = 260 nm for RNA molecules) and electrophoretic assay (run on 1.5% agarose gel) was evaluated. The E.E. was determined as the percentage of siRNA encapsulated into SUVs, calculated subtracting the amount of siRNA present in the supernatant of the centrifuged sample from the total amount of siRNA included in the formulation, to the initial amount of siRNA used. In order to evaluate siRNA‐nanoliposome cytotoxicity and their potential in E2F1 silencing, the complexes were transfected at 200 nM in human colorectal adenocarcinoma cell lines HT29 and in human colon mucosa biopsies, isolated from donors during colonoscopy.

The production process (formulation and manufacturing) is of crucial importance in preparing siRNA‐liposome complexes with the desired shape and size without damaging siRNA's integrity. The shape of liposomes is the main factor affecting carrier's entry in the cellular compartment. Spherical nanoparticle's uptake in mammalian cells was demonstrated to be 3.75–5 times more than rod‐shape nanoparticles, indicating that the carrier's curvature can affect the entry in the plasma membrane [43]. In that regard, spherical SUVs were obtained through the ultrasound‐assisted method adopted which has been successfully used to produce stable siRNA‐SUV complexes on nanometric scale. The achieved siRNA‐liposomes, with a mean diameter size of 38 nm (**Table 1**), are useful for the EPR effect, which involves carrier's extravasation through tumour vascular fenestrations of 50–100‐nm range size. Another important feature is the surface charge of liposomes. Zeta‐potential (*ζ*) was investigated for both unloaded and SUVs encapsulating siRNA samples; the results are presented in **Table 1**. The positive zeta‐potential makes the produced liposomes applicable for the encapsulation of negative siRNA molecules and also promotes the fusion with the negatively charged cell membrane. The *ζ*‐value of the unloaded liposomes (27.90 ± 1.60) appears to be significantly higher than the zeta‐potential of the siE2F1‐1324‐SUV sample (18.02 ± 1.07 mV) suggesting that a strong complexation in addition to the siRNA core encapsulation occurred for the loaded structures.

(w/w) siRNA/total lipid ratio.

described (Section 2.1.1).

40 Liposomes

*3.1.2. Results and discussion*

Encapsulation efficiency (E.E.) for siRNA‐loaded nanoliposomes. Results are expressed as average of three determinations with SD as standard deviation [36].

**Table 1.** Size, PDI and zeta‐potential of unloaded and siE2F1‐1324‐loaded small unilamellar vesicles (SUVs) produced.

Considering the high degradability of siRNA molecules, thus the difficulty in preserving them during all production steps, one of the main goals was to produce liposomes on nanometric size ensuring the integrity of siRNA's molecular structure at the end of the process as well as preserving their biological activity. siRNA sequences were encapsulated with a 100% E.E. showing the efficacy of both the production technique and the formulation adopted. Moreover, electrophoretic studies have showed the high complex stability, which is a very important parameter to take into account due to the high degradability and toxicity of free siRNA molecules. This result indicates the safety of the developed ultrasound‐assisted technique which allows preserving siRNA integrity since no evidence of nucleic acids degradation was visible through the electrophoretic assay.

Regarding the unspecific toxicity of SUVs, results have indicated that siRNA‐nanoliposome complexes are far less toxic than Lipofectamine®2000, a commonly used transfection agent which was also investigated in order to have a comparison with the developed liposomal vesicles. siE2F1‐SUVs were also able to significantly reduce the vitality of the HT29 colon carcinoma cells, thus proving the effectiveness of the complexes and their ability in siE2F1 delivery, which finally down‐regulates cell growth. SUVs were able to enter the cell and release siE2F1 without any toxic effects. Finally, a successful uptake and an E2F1‐silencing effect were also observed in cultured human colon mucosa biopsy, achieving an E2F1 protein inhibition till 80.5%, with a patient‐dependent response. It can be stated that the size reduction process through sonication in duty cycle is a far less complex and more rapid method for liposome size reduction than the one usually adopted and is able to produce liposomes in the nanometric size range (which can thus take advantage of the EPR effect) with high degree of size homogeneity and 100% encapsulation efficiency, relevant feature that can guarantee a uniform behaviour in terms of delivery properties [36].

#### **3.2. Nutraceutical application: nanoliposome vectors for ferrous sulphate delivery**

Anaemia, caused by iron deficiencies, is one of the most widespread nutritional deficiencies, affecting globally two billion people [44]. Despite the success of iron food fortification, particularly in developing countries, the lack of a robust, simple and easy‐to‐transfer fortification technology has limited this technology [45]. Moreover, the supplementary micronutrient products present on market in the form of tablet or capsules have to be improved in quality and variety in order to increase their availability and access in the commercial sector [46]. In order to meliorate the supplementary iron products currently on the market, often composed by micrometric particles, sometimes containing the less absorbable ferric form of iron and obtained, in the most of the cases, by using ineffective production processes and drastic conditions, ferrous sulphate nanoliposomes were produced by using the developed simil‐microfluidic apparatus.

#### *3.2.1. Materials and methods*

*Formulation*: l‐α‐phosphatidylcholine from soybean, Type II‐S, 14–23% choline basis (CAS n. 8002‐43‐5), cholesterol (CAS n. 57‐88‐5), ferrous sulphate heptahydrate (CAS n. 7782‐63‐0), ascorbic acid (CAS n. 50‐81‐7) and ethanol of analytical grade (CAS n. 64‐17‐5) were purchased from Sigma‐Aldrich (Milan, Italy) and used for liposome production. Unloaded and ferrous sulphate‐loaded nanoliposomes were produced by using the simil‐microfluidic bench‐scale apparatus developed whose layout and main process steps are described in Section 2.2.1. In particular, a 10:1 (Vhs/Vls) volumetric flow rate ratio and a 5‐mg/ml lipid concentration in the final hydroalcoholic solution were used for liposome preparation.

*Production:* Briefly, a lipid/ethanol solution was obtained by dissolving PC and cholesterol in 10 ml of ethanol. Cholesterol was used at 2.5:1 (mol/mol) PC/CHOL ratio which corresponds to the typical composition of the cell membrane, as suggested by Abbasi and Azari [47] and was added to the formulation in order to stabilize the loaded vesicles. Ferrous sulphate heptahydrate and ascorbic acid were dissolved in 100 ml of deionized water, which was used as hydration solution. Ascorbic acid was added as an anti‐oxidant to preserve the ferrous ion against oxidation in a ferrous/ascorbic acid. It has been shown that the co‐addition of ascorbic acid and iron in a 2:1 molar ratio (6:1 weight ratio) increases iron absorption from foods twofold to threefold in adults and children [48–50]. In particular, different formulations were produced which differ from each other for the ferrous sulphate/total formulation components (lipids, ascorbic acids and ferrous sulphate) weight ratio (w/w) used. Starting from a 0.06 ferrous sulphate/total components weight ratio, selected from Xia and Xu [51], nanoliposomes were also produced by using a 0.02 ferrous sulphate/total components (w/w) ratio and maintaining all the other chemical and adopted process parameters constant. In order to have a comparison with the ferrous sulphate‐loaded particles, unloaded nanoliposomes were produced by adoperating the same formulation and process conditions but using, as hydration solution, pure deionized water without the addition of ascorbic acid and iron.

Unloaded and ferrous sulphate‐loaded SUVs were at first characterized in terms of morphology, size and zeta‐potential (ZetasizerNano ZS, Malvern, UK). The resulting particle size distribution was plotted as the number of liposomes versus size.

The encapsulation efficiency was determined as the percentage of ferrous sulphate encapsulated in nanoliposomes to the initial amount of ferrous sulphate included in the formulation. Triton X100 at 1% (v/v) was used in order to lyse the nanoliposomes and analyse the encapsulated ferrous sulphate. Iron determination was performed by the 1,10‐phenanthroline colorimetric method through UV spectrophotometric assay (Lambda 25 UV/VIS Spectrophotometer, Perkin Elmer, Monza, Italy). A *λ* = 510 nm, typical of the 1,10‐phenanthroline‐Fe2+ ions complex, was considered.

#### *3.2.2. Results and discussion*

sector [46]. In order to meliorate the supplementary iron products currently on the market, often composed by micrometric particles, sometimes containing the less absorbable ferric form of iron and obtained, in the most of the cases, by using ineffective production processes and drastic conditions, ferrous sulphate nanoliposomes were produced by using the

*Formulation*: l‐α‐phosphatidylcholine from soybean, Type II‐S, 14–23% choline basis (CAS n. 8002‐43‐5), cholesterol (CAS n. 57‐88‐5), ferrous sulphate heptahydrate (CAS n. 7782‐63‐0), ascorbic acid (CAS n. 50‐81‐7) and ethanol of analytical grade (CAS n. 64‐17‐5) were purchased from Sigma‐Aldrich (Milan, Italy) and used for liposome production. Unloaded and ferrous sulphate‐loaded nanoliposomes were produced by using the simil‐microfluidic bench‐scale apparatus developed whose layout and main process steps are described in Section 2.2.1. In particular, a 10:1 (Vhs/Vls) volumetric flow rate ratio and a 5‐mg/ml lipid concentration in the

*Production:* Briefly, a lipid/ethanol solution was obtained by dissolving PC and cholesterol in 10 ml of ethanol. Cholesterol was used at 2.5:1 (mol/mol) PC/CHOL ratio which corresponds to the typical composition of the cell membrane, as suggested by Abbasi and Azari [47] and was added to the formulation in order to stabilize the loaded vesicles. Ferrous sulphate heptahydrate and ascorbic acid were dissolved in 100 ml of deionized water, which was used as hydration solution. Ascorbic acid was added as an anti‐oxidant to preserve the ferrous ion against oxidation in a ferrous/ascorbic acid. It has been shown that the co‐addition of ascorbic acid and iron in a 2:1 molar ratio (6:1 weight ratio) increases iron absorption from foods twofold to threefold in adults and children [48–50]. In particular, different formulations were produced which differ from each other for the ferrous sulphate/total formulation components (lipids, ascorbic acids and ferrous sulphate) weight ratio (w/w) used. Starting from a 0.06 ferrous sulphate/total components weight ratio, selected from Xia and Xu [51], nanoliposomes were also produced by using a 0.02 ferrous sulphate/total components (w/w) ratio and maintaining all the other chemical and adopted process parameters constant. In order to have a comparison with the ferrous sulphate‐loaded particles, unloaded nanoliposomes were produced by adoperating the same formulation and process conditions but using, as hydration

final hydroalcoholic solution were used for liposome preparation.

solution, pure deionized water without the addition of ascorbic acid and iron.

tribution was plotted as the number of liposomes versus size.

plex, was considered.

Unloaded and ferrous sulphate‐loaded SUVs were at first characterized in terms of morphology, size and zeta‐potential (ZetasizerNano ZS, Malvern, UK). The resulting particle size dis-

The encapsulation efficiency was determined as the percentage of ferrous sulphate encapsulated in nanoliposomes to the initial amount of ferrous sulphate included in the formulation. Triton X100 at 1% (v/v) was used in order to lyse the nanoliposomes and analyse the encapsulated ferrous sulphate. Iron determination was performed by the 1,10‐phenanthroline colorimetric method through UV spectrophotometric assay (Lambda 25 UV/VIS Spectrophotometer, Perkin Elmer, Monza, Italy). A *λ* = 510 nm, typical of the 1,10‐phenanthroline‐Fe2+ ions com-

developed simil‐microfluidic apparatus.

*3.2.1. Materials and methods*

42 Liposomes

The simil‐microfluidic bench‐scale apparatus developed has allowed to successfully produce iron‐loaded nanoliposomes in the desired range size. The method has permitted to produce ferrous sulphate‐nanostructured vectors without the use of drastic conditions, such as solvents and/or high pressure, currently used in literature and also at industrial scale for iron particle manufacturing by using discontinuous and laborious processes such as reverse phase evaporation, thin‐film hydration and homogenization freeze‐thawing production methods [47, 51–53]. A part of the drastic conditions used, limits in the output volumes of final product, usually ranging from 10 to 60 mL with the above‐mentioned techniques, represents another crucial problems directly linked with high commercial costs of supplemental products which, for this reason, are not yet widely used as very proper therapies. With the simil‐microfluidic setup developed, by using the ultrasound as tool for the process intensification, it was possible to obtain a massive output with the minimum energy, costs and time by operating in a semi‐continuous manner. In particular, spherical liposomes were obtained in a nanometric range size as shown in **Table 2**.

Taking into account that the iron solubility is very dependent on the size and the shape of the iron particle complexes, characteristics which are governed by the manufacturing process [45], the simil‐microfluidic setup realized was successfully applied for ferrous sulphate nanoliposomes production. In that regard, particles on nanometric scale are required to maintain the transparency of clear beverages during enrichment: carriers have to be small enough so as not to scatter light and be detected by naked eye [54].

The nanoscale plays a crucial role also for other forms of iron supplementation such as oral formulations, the first choice to replace normal iron levels. In this case, the size of nanoparticle systems has a remarkable influence on carrier's uptake after their administration: in many works, it has been proven that nanostructured delivery systems yield an increase in drug uptake, enhancing the intestinal absorption of the active principle [55, 56]. As shown in **Table 2**, vesicles of 48–65‐nm diameter range size have been successfully obtained through the developed setup with PDI values of 0.38 ′ 0.01 and 0.63 ′ 0.12, respectively, for the 0.06 and 0.02 (w/w) formulations produced. Due to the presence of polyunsaturated fatty acids (linoleic and oleic acids) composing the phosphatidylcholine, vesicles presented negative zeta‐potential values (more negative for the 0.06, w/w formulation due to the presence of large amounts of ascorbic acid which decreases the pH of the sample) and an encapsulation efficiency that increases with

**Table 2.** Size, PDI, zeta‐potential and encapsulation efficiency of loaded small unilamellar vesicles (SUVs) produced, obtained at different weight ratios of ferrous sulphate to the total formulation components.

increasing of the lipid amount with respect to those of iron. In particular, as can be seen in **Table 2**, the encapsulation efficiency increases from about 22 to 52% when the weight ratio between ferrous sulphate and the total lipid decreases from 0.06 to 0.02 (w/w).

### **4. Conclusions**

Based on the use of ultrasound as alternative energy resource, a solid particles size reduction/homogenization process was developed and coupled with the bench‐scale conventional thin‐film hydration method. The technique was developed in order to produce, in a versatile manner, nanometric structures, with the desired dimension.

Moreover, due to its easiness, reliability, versatility and its great potential in reducing time spent, the ultrasound intensification tool was also used for liposome homogenization operation during vesicles production through the developed simil‐microfluidic technique.

The phenomenology involved in liposome formation was described for both the methods; applications regarding the entrapment of active molecules were also described as case histories.

siRNA‐nanoliposome complexes (for gene therapy application) were produced for the inhibition of E2F1 protein expression, studied as a potential way to treat colorectal cancer. By the ultrasound‐assisted thin‐film hydration technique, nanoliposomes with 33–38‐nm range size and 100% siRNA encapsulation efficiency were obtained. The produced loaded SUVs demonstrated a very low cytotoxicity in cells when compared with the commercial transfection agent Lipofectamine®2000 and an excellent uptake in the cultured human colon mucosa tissues. A remarkable effect on anti‐E2F1 expression after a transfection of siE2F1‐1324‐ SUV sample has been demonstrated also in a dynamic human model such the colon tissue microenvironment.

For nutraceutical application, nanoliposomes loaded with ferrous sulphate with good dimensional features (48–65 nm vesicles) and encapsulation efficiency were successfully produced using the developed simil‐microfluidic apparatus, avoiding the use of toxic solvents and drastic conditions.

All the achieved positive results endorse the usefulness of both the formulative and the plant‐ engineering approaches adopted for nanostructured vectors production to be used in pharmaceutical and nutraceutical applications.

## **Author details**

Sabrina Bochicchio1,2, Gaetano Lamberti1 and Anna Angela Barba2 \*

\*Address all correspondence to<EMAIL_ADDRESS>
1 Department of Industrial Engineering, University of Salerno, Fisciano, Salerno, Italy

2 Department of Pharmacy, University of Salerno, Fisciano, Salerno, Italy

## **References**

increasing of the lipid amount with respect to those of iron. In particular, as can be seen in **Table 2**, the encapsulation efficiency increases from about 22 to 52% when the weight ratio

Based on the use of ultrasound as alternative energy resource, a solid particles size reduction/homogenization process was developed and coupled with the bench‐scale conventional thin‐film hydration method. The technique was developed in order to produce, in a versatile

Moreover, due to its easiness, reliability, versatility and its great potential in reducing time spent, the ultrasound intensification tool was also used for liposome homogenization opera-

The phenomenology involved in liposome formation was described for both the methods; applications regarding the entrapment of active molecules were also described as case histories. siRNA‐nanoliposome complexes (for gene therapy application) were produced for the inhibition of E2F1 protein expression, studied as a potential way to treat colorectal cancer. By the ultrasound‐assisted thin‐film hydration technique, nanoliposomes with 33–38‐nm range size and 100% siRNA encapsulation efficiency were obtained. The produced loaded SUVs demonstrated a very low cytotoxicity in cells when compared with the commercial transfection agent Lipofectamine®2000 and an excellent uptake in the cultured human colon mucosa tissues. A remarkable effect on anti‐E2F1 expression after a transfection of siE2F1‐1324‐ SUV sample has been demonstrated also in a dynamic human model such the colon tissue

For nutraceutical application, nanoliposomes loaded with ferrous sulphate with good dimensional features (48–65 nm vesicles) and encapsulation efficiency were successfully produced using the developed simil‐microfluidic apparatus, avoiding the use of toxic solvents and dras-

All the achieved positive results endorse the usefulness of both the formulative and the plant‐ engineering approaches adopted for nanostructured vectors production to be used in phar-

1 Department of Industrial Engineering, University of Salerno, Fisciano, Salerno, Italy

2 Department of Pharmacy, University of Salerno, Fisciano, Salerno, Italy

and Anna Angela Barba2

\*

tion during vesicles production through the developed simil‐microfluidic technique.

between ferrous sulphate and the total lipid decreases from 0.06 to 0.02 (w/w).

manner, nanometric structures, with the desired dimension.

**4. Conclusions**

44 Liposomes

microenvironment.

tic conditions.

**Author details**

maceutical and nutraceutical applications.

Sabrina Bochicchio1,2, Gaetano Lamberti1

\*Address all correspondence to<EMAIL_ADDRESS>

[32] Lasic DD, Papahadjopoulos D. Medical Applications of Liposomes: Elsevier; 1998, Amsterdam, The Netherlands

[16] Bangham A, Horne R. Negative staining of phospholipids and their structural modification by surface‐active agents as observed in the electron microscope. Journal of

[17] Wagner A, Vorauer‐Uhl K. Liposome technology for industrial purposes. Journal of

[18] Jaafar‐Maalej, Chiraz, Catherine Charcosset, and Hatem Fessi. A new method for liposome preparation using a membrane contactor. Journal of Liposome Research.

[19] Sekhon BS. Supercritical fluid technology: an overview of pharmaceutical applications.

[20] Jahn A, Vreeland WN, DeVoe DL, Locascio LE, Gaitan M. Microfluidic directed forma-

[21] Pons M, Foradada M, Estelrich J. Liposomes obtained by the ethanol injection method.

[22] Yu B, Lee RJ, Lee LJ. Microfluidic methods for production of liposomes. Methods in

[23] Avvaru B, Patil MN, Gogate PR, Pandit AB. Ultrasonic atomization: effect of liquid phase

[24] Kull H‐J. Theory of the Rayleigh‐Taylor instability. Physics Reports. 1991;**206**(5):197‐325 [25] Abramov OV. High‐intensity Ultrasonics: Theory and Industrial Applications: CRCPress;

[26] Huang X, Caddell R, Yu B, Xu S, Theobald B, Lee LJ, et al. Ultrasound‐enhanced micro-

[27] Woodbury DJ, Richardson ES, Grigg AW, Welling RD, Knudson BH. Reducing liposome size with ultrasound: bimodal size distributions. Journal of Liposome Research.

[28] Barba A, Bochicchio S, Lamberti G, Dalmoro A. Ultrasonic energy in liposome production: process modelling and size calculation. Soft Matter. 2014;**10**(15):2574‐81

[29] Helfrich W. The size of bilayer vesicles generated by sonication. Physics Letters A.

[30] Helfrich W. Elastic properties of lipid bilayers: theory and possible experiments. Zeitschrift für Naturforschung Teil C: Biochemie, Biophysik, Biologie, Virologie.

[31] Fromherz P. Lipid‐vesicle structure: size control by edge‐active agents. Chemical Physics

fluidic synthesis of liposomes. Anticancer Research. 2010;**30**(2):463‐6

International Journal of PharmTech Research, 2010;**2**(1):810‐26

International Journal of Pharmaceutics. 1993;**95**(1‐3):51‐6

tion of liposomes of controlled size. Langmuir. 2007;**23**(11):6289‐93

Molecular Biology. 1964;**8**(5):660‐IN10

Drug Delivery. 2010;**2011**

Enzymology. 2009;**465**:129-41

properties. Ultrasonics. 2006;**44**(2):146‐58

1999, Amsterdam, The Netherlands

2006;**16**(1):57‐80

1974;**50**(2):115‐6

1973;**28**(11):693

Letters. 1983;**94**(3):259‐66

2011;**21**(3):213‐20

46 Liposomes

## **Chapter 3**

**Provisional chapter**

## **Lipobeads**

**Lipobeads**

[45] Mehansho H. Iron fortification technology development: new approaches. The Journal

[46] Mora JO. Iron supplementation: overcoming technical and practical barriers. The Journal

[47] Abbasi S, Azari S. Efficiency of novel iron microencapsulation techniques: fortification of milk. International Journal of Food Science & Technology. 2011;**46**(9):1927‐33

[48] Stekel A, Monckeberg F, Beyda V. Combating iron deficiency in Chile: a case study: International Life Sciences Institute‐Nutrition Foundation, 1986, Washington, D.C., USA

[49] Stekel A, Olivares M, Pizarro F, Chadud P, Lopez I, Amar M. Absorption of fortification iron from milk formulas in infants. The American Journal of Clinical Nutrition.

[50] Lynch SR, Stoltzfus RJ. Iron and ascorbic acid: proposed fortification levels and recom-

[51] Xia S, Xu S. Ferrous sulphate liposomes: preparation, stability and application in fluid

[52] Kosaraju SL, Tran C, Lawrence A. Liposomal delivery systems for encapsulation of ferrous sulphate: preparation and characterization. Journal of Liposome Research.

[53] De Paoli T, Hager AA. Liposomes containing bioavailable iron [II] and process for

[54] Danino D, Livney YD, Ramon O, Portnoy I, Cogan U. Beta‐casein assemblies for enrichment of food and beverages and methods of preparation thereof. Google Patents; 2014

[55] Desai MP, Labhasetwar V, Amidon GL, Levy RJ. Gastrointestinal uptake of biodegradable microparticles: effect of particle size. Pharmaceutical Research. 1996;**13**(12):1838‐45

[56] Hussain N, Jaitley V, Florence AT. Recent advances in the understanding of uptake of microparticulates across the gastrointestinal lymphatics. Advanced Drug Delivery

mended iron compounds. The Journal of Nutrition. 2003;**133**(9):2978S‐84S

milk. Food Research International. 2005;**38**(3):289‐96

obtaining them. Google Patents; 1996

Reviews. 2001;**50**(1):107‐42

of Nutrition. 2006;**136**(4):1059‐63

of Nutrition. 2002;**132**(4):853S‐5S

1986;**43**(6):917‐22

48 Liposomes

2006;**16**(4):347‐58

#### Sergey Kazakov Additional information is available at the end of the chapter

Sergey Kazakov

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/intechopen.70056

#### **Abstract**

A combination of lipid bilayer and cross-linked polymer network is the logical step in development of polymeric and liposomal nanoscopic systems to provide the natural level of functionality. From liposomal systems, lipobeads borrow the welldeveloped methods for preparation, diversity of lipids to control the properties of a lipid bilayer, biocompatibility of the lipid bilayer, possibility to vary size and morphology (passive targeting), availability of the external surface for attachment of various ligands (active targeting), encapsulation efficiency of both hydrophilic and hydrophobic molecules. Mechanical stability of the lipid bilayer and environmental responsiveness of the whole structure are the properties that hydrogel core brings to the new construct. The reports reviewed in this chapter demonstrate that lipobeads of nanometer and micrometer sizes can be prepared in different media, retain their stimuli responsiveness under physiological conditions, exhibit both reversible and irreversible aggregation, can be loaded with both small and high molecular weight molecules. As a platform for drug delivery systems, lipobeads have already been loaded with chemotherapeutics, malaria vaccine, and dermatological agent providing different controlled release profiles without leakage. Consecutive multistep triggering, new schemes of drug release, combined drug delivery, vesobeads, proteolipobeads, enzyme-containing lipobeads, and hemi-lipobeads are the directions for their future development.

DOI: 10.5772/intechopen.70056

**Keywords:** liposomes, nanogels, supramolecular assembly, lipobeads, preparation, properties, application

## **1. Introduction: why lipobeads?**

In accord with our understanding of complex biological mechanisms prevailing *in situ*, the arrangement of a lipid bilayer/hydrogel assembly—the lipid vesicles filled with polymeric

networks (**Figure 1**) achievable experimentally in laboratory—could be a logical step in the development of polymeric and liposomal-beaded nanoscopic systems.

In the last several decades, different terminology, such as supramolecular biovectors (SMBVs), lipid-coated microgels, lipobeads (LB), gel-filled vesicles, lipogels, gel core liposomes, microgel-in-liposomes, hydrogel-supported lipid bilayer, and nanolipogels particles (nLG), has been utilized to describe these lipid membrane/hydrogel structures. In this chapter, we use the term "lipobeads" for the spherical bipartite structures made of a hydrogel core enclosed within a lipid bilayer.

Actually, the bicompartmental structure of lipobeads mimics natural arrangements of living cells. Just look at the cell envelopes for all three main domains of life (*eubacteria*, *archaea*, and *eukaryotes*). They represent a successive organization of the macromolecular networks (cytoplasm, cell wall, capsule, etc.) and lipid bilayers (cell bilayer membranes and internal membrane system), which Nature uses to provide workability, multifunctionality, and dynamism of living cells of different types.

Evidently, the first synthetic lipid vesicles filled with hydrogel (lipobeads) were reported in 1987, when a successful polymerization within liposomes had been accomplished [1] and microspherules of agarose-gelatin gels filled with gold particles had been mixed with liposomes in the course of their preparation [2]. In 1989, a concept of supramolecular biovectors (SMBVs) was filed as a patent application [3]. The SMBV system was prepared from polysaccharide gel fragments obtained by disruption of a gel of chemically cross-linked maltodextrins and subsequently phospholipidated. In 1994, the SMBVs were reported as new carriers of active substances, such as interleukin-2 (IL-2) [4]. In 1995, lipobeads with Ca-alginate hydrogel core were obtained as a byproduct of a method for the preparation of Ca-alginate hydrogel nanoparticles using the internal compartment of liposomes [5]. In 1996, the spherical hydrogel/lipid bilayer constructs for the first time were named as "lipobeads," and it was shown that a lipid bilayer was formed on the surface of hydrogel polymer beads upon the addition of phospholipids, if their surface had been modified with the covalently attached fatty acids [6]. Lipobeads with an environmentally sensitive hydrogel core prepared by hydration of lipid films with microgel suspension were described as drug delivery systems in 1998 [7]. In the early 2000s, photopolymerization within liposomes was used for preparation of the so-called synthetic polymer complements with imprinted recognition sites [8] and the environmentally responsive nanogels [9]. The latter work contributed toward the characterization of the compatibility of nanogels

**Figure 1.** Schematic of a spherical lipid bilayer/hydrogel assembly (lipobead).

and phospholipid bilayer leading to spontaneous phospholipidation of nanogels [10]. Further studies on lipobeads development were devoted either to new methodologies including different compositions of hydrogel core or different agents which could be loaded into the lipobeads.

The chapter has a comprehensive view on (1) synthetic feasibility, functionality and characterization of lipobeads (Sections 2 and 3) and (2) their potential applications as precursors for novel encapsulated and combined drug delivery systems, as microbiochemical reactors, and as an experimental model to study the origin of life (Section 4). It is a useful source of references for the researchers from both academia and industry, who deal with the aforementioned areas of applications. It may be predicted that in the future, the demand in this information will rise dramatically because of a growing interest, especially, in the encapsulated drug delivery systems with tiny bioscopic mechanisms of drug release.

## **2. Strategies of lipobeads preparation**

networks (**Figure 1**) achievable experimentally in laboratory—could be a logical step in the

In the last several decades, different terminology, such as supramolecular biovectors (SMBVs), lipid-coated microgels, lipobeads (LB), gel-filled vesicles, lipogels, gel core liposomes, microgel-in-liposomes, hydrogel-supported lipid bilayer, and nanolipogels particles (nLG), has been utilized to describe these lipid membrane/hydrogel structures. In this chapter, we use the term "lipobeads" for the spherical bipartite structures made of a hydrogel core enclosed

Actually, the bicompartmental structure of lipobeads mimics natural arrangements of living cells. Just look at the cell envelopes for all three main domains of life (*eubacteria*, *archaea*, and *eukaryotes*). They represent a successive organization of the macromolecular networks (cytoplasm, cell wall, capsule, etc.) and lipid bilayers (cell bilayer membranes and internal membrane system), which Nature uses to provide workability, multifunctionality, and dynamism

Evidently, the first synthetic lipid vesicles filled with hydrogel (lipobeads) were reported in 1987, when a successful polymerization within liposomes had been accomplished [1] and microspherules of agarose-gelatin gels filled with gold particles had been mixed with liposomes in the course of their preparation [2]. In 1989, a concept of supramolecular biovectors (SMBVs) was filed as a patent application [3]. The SMBV system was prepared from polysaccharide gel fragments obtained by disruption of a gel of chemically cross-linked maltodextrins and subsequently phospholipidated. In 1994, the SMBVs were reported as new carriers of active substances, such as interleukin-2 (IL-2) [4]. In 1995, lipobeads with Ca-alginate hydrogel core were obtained as a byproduct of a method for the preparation of Ca-alginate hydrogel nanoparticles using the internal compartment of liposomes [5]. In 1996, the spherical hydrogel/lipid bilayer constructs for the first time were named as "lipobeads," and it was shown that a lipid bilayer was formed on the surface of hydrogel polymer beads upon the addition of phospholipids, if their surface had been modified with the covalently attached fatty acids [6]. Lipobeads with an environmentally sensitive hydrogel core prepared by hydration of lipid films with microgel suspension were described as drug delivery systems in 1998 [7]. In the early 2000s, photopolymerization within liposomes was used for preparation of the so-called synthetic polymer complements with imprinted recognition sites [8] and the environmentally responsive nanogels [9]. The latter work contributed toward the characterization of the compatibility of nanogels

development of polymeric and liposomal-beaded nanoscopic systems.

**Figure 1.** Schematic of a spherical lipid bilayer/hydrogel assembly (lipobead).

within a lipid bilayer.

50 Liposomes

of living cells of different types.

Two general methods available to date for preparation of artificial bilayer-coated hydrogel particles (lipobeads, giant, or nano) are sketched in **Figure 2**.

The first one (**Figure 2A**) uses the liposomal interior as a chemical reactor for the formation of hydrogel by polymerization [1, 8–22]. In this method, lipid bilayer should be sealed enough to retain the concentration of pregel components and strong enough to withstand the steps preventing polymerization in the surrounding medium.

The second one (**Figure 2B**) is based on the formation of lipid layers around hydrogels after microgel/liposome mixing. In this case, lipid bilayer adsorption on the surface of hydrogel particles prepared separately was promoted via Coulombic attraction between the charged microgels and oppositely charged lipids [7, 23, 24], hydration of lipid films with micro- or nanogel suspension [2, 3], introduction of hydrophobic anchors at the microgel surface around which adsorbing lipids may assemble [6, 25–28], centrifugation of microgels onto a lipid film [29], microfluidic flowing [21], and emulsification [30–32].

In both cases, it is the stability, fluidity, and permeability that are the main properties of a lipid bilayer, which should be governed in the course of lipobeads engineering.

#### **2.1. Effectors of the lipid bilayer fluidity, stability, and permeability**

Stability and permeability of a lipid bilayer depend on its fluidity, which can vary with temperature and phospholipid composition. Bilayers undergo a change from liquid to gel (solid) state at the so-called lipid (or order-disorder) phase transition temperature (*T*t ) characteristic to a phospholipid used (**Figure 3**).

Therein, the shorter the length of hydrocarbon chains, the lower is the *T*t , a long hydrocarbon chain at sn-1 position and a short chain at sn-2 position on glycerol exhibit a lower *T*t than that of a phospholipid with the opposite arrangement, presence, and position of double bonds in the hydrocarbon chain makes *T*t lower than that of the saturated analogue; a bulky

**Figure 2.** Two methods for lipobeads preparation: (A) formation of hydrogel core by polymerization within vesicle interior and (B) mixing of separately prepared lipid vesicles and hydrogel particles (microgels or nanogels).

**Figure 3.** Structure of lipid bilayer below and above the order-disorder transition temperature.

head group confers the lipid a lower *T*t than it would be with a smaller head group, the phospholipids with negatively charged head groups favor a lower *T*t than that of an uncharged phospholipid. Ionic conditions can modulate this effect, for example, presence of cations in the surrounding water increases *T*t .

In the liquid-crystalline "disordered" state, the membrane is fluidic, namely: (i) both alkane chains and head groups of phospholipids are more flexible than in the solid "ordered" state, (ii) the area lipids occupy becomes greater by changing from a 0.48 nm2 /head group to 0.7 nm2 /head group, that is, bilayer expands, (iii) lateral diffusion of phospholipids in the plane of the bilayer and rotation of lipid molecules around C─C bonds accelerate, and (iv) transverse "flip-flop" migration of lipids from one monolayer to the other side of the bilayer becomes more probable.

The factors affecting the lipid phase transition temperature are crucial for lipobeads engineering. Indeed, one can expect the lipid bilayer to be more elastic (favorable for formation of unilamellar membrane) and less sealed (unfavorable for gelation within liposomal interior) above *T*t than below *T*t . It is known that stability and permeability of naturally occurred membranes can be varied by balancing composition of cholesterol and alcohols. Particularly, cholesterol strongly interacts with phospholipids and inhibits the passive permeability of lipid membrane to water and small electrolytes and nonelectrolytes. The extent of "sealing" directly depends on the amount of cholesterol present, up to moderate levels of cholesterol. However, at very high levels of cholesterol, pure cholesterol phase separates out and leads to an increased leakage through interfacial lipids and unstable aggregates of cholesterol. On the contrary, the insertion of anesthetic molecules, such as alcohols, into the membrane increases the membrane fluidity at a given temperature by depression of the lipid order-disorder transition temperature. Additionally, sphingolipids are commonly believed to protect the cell surface against harmful environmental factors by forming a mechanically stable and chemically resistant outer leaflet of the plasma membrane.

#### **2.2. Polymerization within liposomal nano-/microreactors**

head group confers the lipid a lower *T*t

the surrounding water increases *T*t

52 Liposomes

pholipids with negatively charged head groups favor a lower *T*t

lipids occupy becomes greater by changing from a 0.48 nm2

.

**Figure 3.** Structure of lipid bilayer below and above the order-disorder transition temperature.

phospholipid. Ionic conditions can modulate this effect, for example, presence of cations in

**Figure 2.** Two methods for lipobeads preparation: (A) formation of hydrogel core by polymerization within vesicle

interior and (B) mixing of separately prepared lipid vesicles and hydrogel particles (microgels or nanogels).

In the liquid-crystalline "disordered" state, the membrane is fluidic, namely: (i) both alkane chains and head groups of phospholipids are more flexible than in the solid "ordered" state, (ii) the area

that is, bilayer expands, (iii) lateral diffusion of phospholipids in the plane of the bilayer and rotation of lipid molecules around C─C bonds accelerate, and (iv) transverse "flip-flop" migration of

lipids from one monolayer to the other side of the bilayer becomes more probable.

than it would be with a smaller head group, the phos-

/head group to 0.7 nm2

than that of an uncharged

/head group,

In general, preparation of lipobeads using vesicle interior as microreactors includes a number of crucial steps, as shown in **Figure 2A**.

First of all, encapsulation of hydrogel-forming components into liposomes can result from hydration of lipid cast film formed upon solvent evaporation [1, 2, 5, 9, 11, 13, 14, 16, 32], electroformation [12, 22, 33, 34], or rapid phase evaporation [19, 20]. The size of liposomes ensures the final size of lipobeads. Depending on the size, lipobeads can be classified into two groups: nanolipobeads (<1000 nm) and giant lipobeads (>1 μm). Nanolipobeads should be used in realistic drug delivery systems, whereas giant lipogels permit a direct study of structural functionality of the hydrogel/lipid bilayer assemblies, cargo loading, and release mechanisms [12] using optical microscopy.

For 100-nm lipobeads, the liposomal formulations have to be sonicated [1, 9] or extruded through a nanopore filter of a needed pore size [5, 8, 11, 14, 16, 19, 32]. Another approach to control the size of pre-lipobeads is based on hydrodynamic focusing of a stream of the liposome precursor solution by the flow of a hydrogel-forming solution within a microfluidic device [21]. Although the microfluidic-directed approach and electroformation are very elegant methods, they are hardly suitable for a scaled production of lipobeads with regards to pharmaceutical applications.

If the ultimate goal was to engineer the giant lipobeads greater than 1 μm, the gentle hydration of a phospholipid cake [17, 36] or a hybrid agarose/lipid film [unpublished], electroformation [12, 22, 33, 34], emulsification [32], and reverse phase evaporation [1, 13, 30] were used without any agitation of the lipid formulations.

Typical lipid formulations for preparation of lipid vesicles filled with the hydrogel-forming solution are shown in **Table 1**.

**Table 1.** Lipid formulations used for preparation of lipobeads by polymerization within lipid vesicles.

Second, when a suspension of vesicles filled with the hydrogel-forming solution is prepared, it is important to prevent cross-linking or polymerization outside those vesicles. This has been done by several methods, such as a 5- to 20-fold dilution [9, 11, 12, 14, 16, 22, 33, 34], gel filtration [1, 21], centrifugation and dialysis [5, 13, 20, 30], or introduction of polymerization scavengers (e.g., ascorbic acid [19]) into the extravesicular space. In addition, hydrogel-forming solution as well as cross-linker and initiator can be microinjected directly into the internal compartment of a giant unilamellar phospholipid vesicle (GUV) [17].

The third step is gelation of the hydrogel-forming solution entrapped within the closed lipid bilayer. Thermal and ionotropic cross-linking are the examples of physical cross-linking reactions. Some hydrogel cores were made of polysaccharides when temperature changes [2, 5, 32]. Indeed, agarose [5] and κ-carrageenan [32] are the temperature-sensitive polysaccharides which structure in aqueous solutions undergoes a transition from a random-coil conformation to the cross-linked double helixes upon cooling. Agarose is not biodegradable, but its combination with gelatin can bring biodegradability [38]. Gelatin is a thermoresponsive protein, forming a reversible cross-linked network by cooling a water-based solution of the polymer below 35°C. The hydrogel can be liquefied by heating it to physiological temperatures. Interestingly, κ-carrageenan, an anionic polysaccharide carrying one sulfate group, can be cross-linked both thermally (upon cooling) and ionotropically in the presence of divalent or monovalent cations [38]. Similar to alginate, the degradation of carrageenan hydrogels is driven by the exchange of ions with the surrounding fluids. In the course of ionic cross-linking within interior of vesicles, the sections of the polymer backbone carrying the charge bind with ions of opposite charge. For example, when multivalent cations (e.g., Ca2+) were added to a water-based alginate [5] or poly(ethylene dioxythiophene)/poly(styrene sulfonate) [17] solutions, they bound adjacent polymer chains forming ionic interchain bridges that caused a cross-linking. The pHdriven cross-linking inside liposome was carried out by lowering the pH of aqueous solution of poly(acrylic acid) carrying carboxyl groups [20, 30].

The greatest portion of works on gelation within liposomal reactor used photopolymerization to generate a strong covalently cross-linked hydrogel [1, 9, 11, 14, 16, 22, 33, 34, 39]. The monomer and cross-linker depends on the hydrogel core properties required for different applications, as shown in **Table 2**.

Finally, the formulation has to be washed from unreacted chemicals using centrifugation and/ or dialysis. In the course of this step, the required medium external to the lipobeads can be introduced. For example, lipobeads could be dispersed in distilled water [13, 30, 36], buffers with pH varied from 7.0 to 7.8 [1, 8, 9, 11, 14, 16, 17, 19–21, 30], or aqueous solutions of sucrose [12, 24, 33, 34]. If necessary, the prepared lipobeads can be dried by gentle evaporation in temperature gradient to be stored at 4°C.

#### **2.3. Hydrogel/liposome mixing**

Second, when a suspension of vesicles filled with the hydrogel-forming solution is prepared, it is important to prevent cross-linking or polymerization outside those vesicles. This has been done by several methods, such as a 5- to 20-fold dilution [9, 11, 12, 14, 16, 22, 33, 34], gel

Abbreviations: DCP: dihexadecyl phosphate, DOPC: 1-2 dioleoyl sn-glycero 3-phosphocholine, DOTAP: dioleoyl trimethylammoniumpropane (positively charged), DPPC: 1,2-dipalmotyl-snglycero-3-phosphatidylcholine, DSPE: 1,2-distearoyl-sn-glycero-3-phosphoethanolamine, EPC: Egg chicken l-α-phosphotidylcholine, HSPC: Hydro Soy l-αphosphotidylcholine, PA: phosphatidic acid, PC: phosphocholine, PE: phosphatidylethanolamine, PEG: polyethylene

**Lipid formulation Method for**

EPC −10°C Cholesterol Reverse phase

NH2

DPPC 42°C — Lipid film hydration,

charged

glycol, PI: phosphatidylinositol, SOPC: 1-stearoyl-*sn*-glycero-3-phosphocholine.

SOPC, neutral 6°C DOTAP, positively

Soybean polar lipid extract: PC (45.7%), PE (22.1%), PI (18.4%), PA (6.9%), others (6.9%)

DOPC −22°C — Electroformation 10–30 μm [12]

HSPC 52°C Cholesterol Lipid film hydration 2–200 μm [36]

**Table 1.** Lipid formulations used for preparation of lipobeads by polymerization within lipid vesicles.

Cholesterol/DCP Hydrodynamic

**Main phospholipid** *T***t Other components**

54 Liposomes

**prelipobead preparation**

evaporation, sonication

sonication

evaporation

extrusion

focusing in microfluidic device

extrusion

? – Lipid film hydration 10–50 μm [17]

Lipid film hydration,

— Emulsification 1–40 μm [32]

DOPC/cholesterol ~100 nm [19]

Cholesterol 0.2–1 μm [8] — 100–200 nm [37]

— Lipid film hydration,

— Detergent removal or extrusion

— Lipid film hydration, extrusion

Cholesterol Reverse phase

**Size Ref.**

~600 nm [1]

~150 nm [9]

~200 nm [13]

~1–2 μm [30]

~100 nm [14]

~120 nm [11]

0.2–100 μm [22, 34]

~800 μm [5]

150–300 nm [21]

~350 nm [16]

From the general scheme of hydrogel/liposome mixing (**Figure 2B**), one can conclude that hydrogel particles and liposomes should be prepared separately, therein the final size of lipobeads will be defined by the size of hydrogel particles prepared before mixing with liposomes. It has been demonstrated microscopically [40, 41] that hydrophobic modification of the nanogels is not required for spontaneous formation of the bilayer on their surface. Together with the other studies [42], these findings presume that hydrogel/lipid bilayer is an energetically favorable structure.

*Abbreviations*: ACVA: 4,4′-azobis(4-cyanovaleric acid); BAA: bis-acrylamido acetate; DEAP: 2,2′-diethoxyacetophenone; dex-HEMA: dextran hydroxyethyl methacrylate; GPC: gel permeation chromatography; IC2959: Irgacure 2959; MBA: *N*,*N*-methylenebisacrylamide; PAA: polyacrylic acid; PAAm: polyacrylamide; PEG: polyethylene glycol; PLA: polylactide; PNIPA: poly(*N*-isopropylacrylamide); PSA: sorbitol acrylate; TEMED: *N,N,N′,N′*-tetramethylethylenediamine; TEGDM: tetraethylene glycol dimethacrylate; VI: 1-vinylimidazole.

**Table 2.** Composition and properties of hydrogel core of lipobeads prepared by polymerization within lipid vesicles.

Lipid formulations used for preparation of lipobeads by hydrogel/liposome mixing are systemized in **Table 3**. Phospholipid with different *T*t were employed to prepare conventional liposomes mostly by the lipid film hydration followed by sonication or extrusion in a variety of buffers at pH 7.0–7.6 and deionized water. Nonetheless, the experiments on giant lipobeads show [36, 41] that injection of ethanol solution of phospholipid into hot water is a promising method for preparation of lipidic formulations, which may allow one to exclude the timeconsuming steps of lipid film formation and hydration and reduce the time for the scaled fabrication of lipobeads from days to hours.

There are only a few reports (**Table 4**) that deal with nanogels to prepare lipobeads on the nanometer scale: one group employed a high pressure homogenizer to crash bulk polysaccharide hydrogel down to nanosized particles [39, 42], the other group used nanogels extracted from liposomal reactors [9, 10, 26, 27]. In principle, emulsion polymerization

*Abbreviations*: DMPC: 1,2-dimyristoyl-snglycero-3-phosphatidylcholine, DMPE: 1,2-Dimyristoyl-sn-glycero-3 phosphoethanolamine, DOPA: dioleoyl glycerol phosphate (negatively charged), DOPC: 1-2 dioleoyl *sn*-glycero 3-phosphocholine, DOPE: dioleoyl glycerol phosphoethanolamine (neutral), DOPG: dioleoylphosphatidylglycerol, DOTAP: dioleoyl trimethylammoniumpropane (positively charged), DPPC: 1,2-dipalmotyl-snglycero-3 phosphatidylcholine, EPC: Egg chicken l-α-phosphotidylcholine, HSPC: Hydro Soy l-α-phosphotidylcholine, PE: phosphatidylethanolamine, PL: phospholipid, POPC: 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, PS – phosphatidylserine, SOPC: 1-stearoyl-*sn*-glycero-3-phosphocholine.

**Table 3.** Lipid formulations used for preparation of lipobeads by hydrogel/liposome mixing.

Lipid formulations used for preparation of lipobeads by hydrogel/liposome mixing are sys-

**Table 2.** Composition and properties of hydrogel core of lipobeads prepared by polymerization within lipid vesicles.

*Abbreviations*: ACVA: 4,4′-azobis(4-cyanovaleric acid); BAA: bis-acrylamido acetate; DEAP: 2,2′-diethoxyacetophenone; dex-HEMA: dextran hydroxyethyl methacrylate; GPC: gel permeation chromatography; IC2959: Irgacure 2959; MBA: *N*,*N*-methylenebisacrylamide; PAA: polyacrylic acid; PAAm: polyacrylamide; PEG: polyethylene glycol; PLA: polylactide; PNIPA: poly(*N*-isopropylacrylamide); PSA: sorbitol acrylate; TEMED: *N,N,N′,N′*-tetramethylethylenediamine; TEGDM:

liposomes mostly by the lipid film hydration followed by sonication or extrusion in a variety of buffers at pH 7.0–7.6 and deionized water. Nonetheless, the experiments on giant lipobeads show [36, 41] that injection of ethanol solution of phospholipid into hot water is a promising method for preparation of lipidic formulations, which may allow one to exclude the timeconsuming steps of lipid film formation and hydration and reduce the time for the scaled

There are only a few reports (**Table 4**) that deal with nanogels to prepare lipobeads on the nanometer scale: one group employed a high pressure homogenizer to crash bulk polysaccharide hydrogel down to nanosized particles [39, 42], the other group used nanogels extracted from liposomal reactors [9, 10, 26, 27]. In principle, emulsion polymerization

were employed to prepare conventional

temized in **Table 3**. Phospholipid with different *T*t

tetraethylene glycol dimethacrylate; VI: 1-vinylimidazole.

PAA MBA IC2959 Radical

PAAm MBA DEAP Dilution 20-fold

**Monomer. Cross-linker Initiator Prevention of**

Anchored and nonanchored PNIPA-VI

56 Liposomes

**macroscale gelation**

scavenging by

by glucose solution (2.8 M)

AA

MBA DEAP Dilution 25-fold Temperature and pH

PAAm MBA ACVA + TEMED GPC Not specified [1] PSA BAA ACVA GPC Not specified [8]

PNIPA or PAAm MBA DEAP Dilution 20-fold Temperature sensitive [14]

Anchored PNIPA TEGDM DEAP Dialysis Temperature

PNIPA MBA DEAP Gel filtration Temperature

dex-HEMA HEMA IC2959 Dilution 10-fold Enzymatically

PNIPA MBA DEAP Dilution Temperature, pH, pI

PLA-PEG-PLA Diacrylate IC2959 Dilution 5-fold Biodegradable [11]

**Property of hydrogel**

sensitivity (probably)

**Ref.**

[9]

[13]

[21]

[16]

[19]

[37]

[22, 33, 34]

**core**

sensitivity

sensitivity

degradable (dextranase)

pH sensitivity (probably)

sensitivity

Enzymes entrapment, storage, protection, and release

fabrication of lipobeads from days to hours.

enables preparation of hydrogel particles with a diameter less than 150 nm. However, there is a problem of complete removal of the residual materials. In the absence of an added surfactant, the method is called precipitation polymerization. With the latter two methods, the lipobeads of 1-μm diameter are produced [2, 7, 23, 25, 28, 31]. To prepare giant lipobeads with a diameter up to 100 μm, the inverse suspension polymerization (ISP) method is commonly applied [6, 24, 35, 43–45].

Liposomes can be brought into contact with hydrogel particles by mixing hydrogel particles and liposomes, addition of hydrogel particles into dried lipid film before hydration or

*Abbreviations*: AA: ascorbic acid, AMPS: 2-acrylamido-2-methyl-1-propanesulfonic acid, dex-HEMA: dextran hydroxyethyl methacrylate, DMAEMA: dimethylaminoethyl methacrylate, E-BIS: *N,N*′-ethylene-bis(acrylamide), ISP: inverse suspension polymerization, MAA: methacrylic acid, MBA: *N*,*N*-methylenebisacrylamide, NPMA: 4-nitrophenyl methacrylate, PAAm: polyacrylamide, PDMAA: polydimethylacrylamide, PEGDA: poly(ethylene glycol) diglycidyl ether, PMAA: polymethacrylic acid, PNIPA – poly(*N*-isopropylacrylamide), PVA: polyvinyl alcohol.

**Table 4.** Composition and properties of hydrogel particles for lipobeads prepared by hydrogel/liposome mixing.

hydration of the lipid film by aqueous suspension of hydrogel particles. The formation of lipid bilayer around hydrogel particles can be enhanced by shaking, vortexing, pipette agitation, centrifugation, freezing-thawing, heating-cooling, or their combination. The fusion of liposomes will be more advanced at temperatures higher than the *T*t of the phospholipid used. Moreover, depending on the electrostatic interaction between bilayer and hydrogel, the liposomes can adsorb on the particles surface, diffuse inside, or/and fuse on the surface with formation of lipobeads [24]. Usually, free liposomes are washed out by centrifugation or removed by ultrafiltration or dialysis. Finally, the lipobeads can be dispersed in a buffer with pH ranged from 7 to 8 or distilled water.

#### **2.4. Modification of hydrogel core and lipid bilayer**

To the great extent, the major methods for lipobeads' synthesis (polymerization within liposomal interior and liposome/hydrogel mixing) and further functionalization are analogous to those used for engineering their compartments—conventional nanogels and lipid bilayers.

#### *2.4.1. Hydrogel core*

hydration of the lipid film by aqueous suspension of hydrogel particles. The formation of lipid bilayer around hydrogel particles can be enhanced by shaking, vortexing, pipette agitation, centrifugation, freezing-thawing, heating-cooling, or their combination. The fusion of

**Table 4.** Composition and properties of hydrogel particles for lipobeads prepared by hydrogel/liposome mixing.

*Abbreviations*: AA: ascorbic acid, AMPS: 2-acrylamido-2-methyl-1-propanesulfonic acid, dex-HEMA: dextran hydroxyethyl methacrylate, DMAEMA: dimethylaminoethyl methacrylate, E-BIS: *N,N*′-ethylene-bis(acrylamide), ISP: inverse suspension polymerization, MAA: methacrylic acid, MBA: *N*,*N*-methylenebisacrylamide, NPMA: 4-nitrophenyl methacrylate, PAAm: polyacrylamide, PDMAA: polydimethylacrylamide, PEGDA: poly(ethylene glycol) diglycidyl ether,

**Hydrogel bead composition Method of bead preparation Size Property of hydrogel beads Ref.**

ISP 5–600 μm –

ISP ~80–100

Sodium hyaluronate-PEGDA Emulsification 1–15 μm Drying/wetting [31]

Extrusion in high-pressure

homogenizer

Precipitation polymerization

Emulsification ~1–5 μm Degradable [2]

ISP 1–100 μm – [46]

ISP ~6 μm pH sensitivity [7]

Emulsification 2–5 μm Degradable [23]

μm

ISP 1–100 μm Electrostatic liposomes/MG

30–60 nm Temperature, pH, pI: core/ bilayer interactions

> Antibody-antigen interaction [45]

– [46]

~0.3–2 μm Temperature, pH sensitivity [25, 28]

interactions

[39]

[35, 43–45]

[24]

used. Moreover, depending on the electrostatic interaction between bilayer and hydrogel, the liposomes can adsorb on the particles surface, diffuse inside, or/and fuse on the surface with formation of lipobeads [24]. Usually, free liposomes are washed out by centrifugation or removed by ultrafiltration or dialysis. Finally, the lipobeads can be dispersed in a buffer with

To the great extent, the major methods for lipobeads' synthesis (polymerization within liposomal interior and liposome/hydrogel mixing) and further functionalization are analogous to those used for engineering their compartments—conventional nanogels and lipid bilayers.

of the phospholipid

liposomes will be more advanced at temperatures higher than the *T*t

PMAA: polymethacrylic acid, PNIPA – poly(*N*-isopropylacrylamide), PVA: polyvinyl alcohol.

pH ranged from 7 to 8 or distilled water.

Agarose-gelatin cross-linked by

Cross-linked polysaccharide

Acylated PVA cross-linked by

PMAA-NPMA cross-linked by

Anchored PDMAA cross-linked

dex-HEMA-MAA (−) or dex-HEMA-DMAEMA (+)

Acylated PVA cross-linked by

Anchored PNIPA/P(NIPA-AA) core-shell cross-linked by MBA

PAAm-allylamine (+) or PAAm-AMPS (−) cross-linked by MBA

cooling

58 Liposomes

fragments

freeze-thaw

MBA

by E-BIS

freeze-thaw

**2.4. Modification of hydrogel core and lipid bilayer**

Polymeric nanogels can be synthesized by three straightforward methods: (i) cross-linking polymer chains within already formed nanoparticles using, for example, emulsion polymerization technique [47–49], (ii) polymerization within the liposomal interior followed by the lipid bilayer removal [9], and (iii) photolithographic fabrication of submicrometer hydrogel particles using the PRINT technique [50, 51] or step and flash imprint lithography (S-FIL) [52] as an alternative nanoimprint photolithographic approach.

To engineer the stimuli-responsive nanogels (**Figure 4**), a molecule of interest can be conjugated to the polymer network through a cleavable tether, so that when the tether is cleaved, the drug is allowed to diffuse into the nearby medium. Alternatively, if different molecules are trapped within an environmentally sensitive polymer network with or without environmentally responsive cleavable linkers, the network either changes its volume (swells/shrinks) or degrades when the environmental conditions change, allowing the molecules to be released. For example [53], the doxorubicin-loaded, pH- and redox-sensitive poly(oligo(ethylene glycol) methacrylates-ss-acrylic acid) nanogels exhibited strong internalization by human hepatocellular carcinoma cells (Bel7402) under reduced opsonization and phagocytosis. Herein, the intracellular glutathione (GSH) triggered the release of doxorubicin from the nanogels into cytosol for subsequent entering the nucleus.

**Figure 4.** A futuristic view on a stimuli-responsive nanogel with entrapped or/and tethered molecules of interest.

#### *2.4.2. Lipid bilayer*

Specific functionality and entrapment of both hydrophilic and hydrophobic molecules into a liposomal interior are the two main objectives of the liposomal system development. Eventually, four types of liposomal systems can be distinguished (**Figure 5**), namely: classical "plain" liposomes, "stealth" liposomes, ligand conjugated liposomes, and stimuli-sensitive liposomes.

The studies on "plain" (traditional) liposomes revealed the difficulty in loading of some types of molecules and leakage of contents from the liposomal interior [54–58]. The further development of the "plain" liposome systems aimed at overcoming these obstacles. In particular, to reduce leakage from liposomes, phospholipids with a higher phase transition temperature [59] were used, and cholesterol [60] and sphingomyelin [61] were incorporated into the lipid bilayer to make it more solid at temperatures of application.

Loading and retention of molecules of interest within liposomes are the molecule dependent processes. For example, weak bases were loaded in response to pH gradients [62–66]. Some molecules, such as doxorubicin, exhibited good retention properties under conditions enhancing their precipitation inside liposomes [67–69], whereas retention of highly hydrophobic molecules, like paclitaxel, was still a challenge [70, 71] until they were converted into the weak bases [72].

Furthermore, the pharmaceutical studies revealed that (i) serum proteins effected on release of drug molecules entrapped into liposomes, (ii) liposomes were cleared very rapidly from

**Figure 5.** Four evolutionary steps of lipid bilayer modification and functionalization.

circulation by uptake into the cells of the mononuclear phagocyte system (MPS), predominantly in the liver and spleen [73, 74], and (iii) there existed both cellular and intracellular barriers to liposomal delivery [75]. The so-called "stealth" liposomes were developed by stabilizing liposomes with protective polymers (e.g., polyethylene glycol, PEG) [76] in order to increase their circulation time within a biologically active environment, such as blood. In addition, the clearance of liposomes from the body was found [74, 80] to be slower, if they contained neutral or slightly negative phospholipids and their size was around 100 nm. Therefore, the rate of molecules' release should be optimized [77–79].

Ligand-conjugated liposomes were built to target specific cells, intracellular organelles, tumor microenvironment, and/or facilitating receptor-mediated endocytosis (attachment of antibodies, folate, transferrin, tyrosine kinase, vascular endothelial growth factor, introduction of fusogenic lipids, and membrane-active peptides). In particular, three ways to facilitate the intracellular drug delivery include (i) introduction of fusogenic lipids or membrane active peptides into liposomal bilayer enhances fusion or even disruption of cell/organelle membrane and thereby improves cytoplasmic delivery of drug [81–85], (ii) utilization of macrophages for natural endocytosis of drug-loaded liposomes [86], and (iii) receptor-mediated endocytosis of ligand-targeted liposomal drug carriers into the intracellular compartment (see reviews [87–90]).

The release of liposomal contents can be triggered either remotely by heat, radiation, and ultrasound or locally by pH, enzymes, and redox triggers (see review [91] and references therein). For these purposes, the lipid bilayer can be modified with stimuli-sensitive phospholipids, polymers, cleavable tethers, and linkers, as shown in **Figure 5**. Recently, the lipid bilayer consisting of molecules of the temperature-sensitive phospholipid 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine (MSPC) and decorated with gold nanorods was shown to become more permeable for the pain blocking molecules (tetrodotoxin) without a tissue burn when exposed repeatedly to a low intensity near-infrared irradiance [92].

#### **2.5. Encapsulation strategies**

*2.4.2. Lipid bilayer*

60 Liposomes

Specific functionality and entrapment of both hydrophilic and hydrophobic molecules into a liposomal interior are the two main objectives of the liposomal system development. Eventually, four types of liposomal systems can be distinguished (**Figure 5**), namely: classical "plain" liposomes, "stealth" liposomes, ligand conjugated liposomes, and stimuli-sensitive liposomes.

The studies on "plain" (traditional) liposomes revealed the difficulty in loading of some types of molecules and leakage of contents from the liposomal interior [54–58]. The further development of the "plain" liposome systems aimed at overcoming these obstacles. In particular, to reduce leakage from liposomes, phospholipids with a higher phase transition temperature [59] were used, and cholesterol [60] and sphingomyelin [61] were incorporated into the lipid

Loading and retention of molecules of interest within liposomes are the molecule dependent processes. For example, weak bases were loaded in response to pH gradients [62–66]. Some molecules, such as doxorubicin, exhibited good retention properties under conditions enhancing their precipitation inside liposomes [67–69], whereas retention of highly hydrophobic molecules, like paclitaxel, was still a challenge [70, 71] until they were converted into the weak bases [72].

Furthermore, the pharmaceutical studies revealed that (i) serum proteins effected on release of drug molecules entrapped into liposomes, (ii) liposomes were cleared very rapidly from

bilayer to make it more solid at temperatures of application.

**Figure 5.** Four evolutionary steps of lipid bilayer modification and functionalization.

#### *2.5.1. Loading during gelation within liposomal interior*

Unfortunately, little work has been done so far to test loading capability of lipobeads prepared by polymerization within the lipid vesicles [11, 14, 15, 20, 30]. Typically, a hydrophobic cargo was either incorporated within lipid bilayer at the step of lipid vesicle formation or copolymerized with hydrogel core as an anchor, whereas a hydrophilic cargo was added as a component of the hydrogel-forming solution and incorporated into the intravesicular space before gelation started [9, 11, 13].

The main challenge of the scheme when a load is introduced into the aqueous phase followed by rehydration of a lipid film and further polymerization within a liposomal reactor might be the damage to the loading molecules by toxic ingredients of the hydrogel-forming solution (if any) and/or high temperature and UV radiation initiating polymerization. This approach can be especially problematic for encapsulation of proteins, because of denaturation. Nonetheless, it has been reported that antigen model (BSA) [30] or combination of protein antigen (Pfs25) and oligonucleotide sequence (CpGODN) encapsulated into pH-cross-linked PAA hydrogel core of lipobeads remain intact and active [20]. Moreover, encapsulation efficiencies of lipobeads were shown to be by 10% higher than those for liposomal carriers. A high encapsulation efficiency of lipobeads has been demonstrated also for hemoglobin [14, 15], which withstands the conditions of free radical polymerization and UV radiation. The other example of successful coencapsulation of hydrophilic proteins (IL-2) and small hydrophobic molecules (TGF-receptor-I inhibitor, SB505124) into the biodegradable hydrogel core of lipobeads has been presented in [11]: encapsulation efficiencies were 80 and 36%, respectively, and UV polymerization did not compromise bioactivity of both immunomodulators. Recently, PAAm lipobeads with a good encapsulation efficiency (37%) of enzymes (bovine Cu, Zn-superoxide dismutase, and bovine milk lactoperoxidase) were synthesized to prove that the UV irradiation used for interior gelation did not cause any reduction in the enzymatic activity of the proteins [37].

#### *2.5.2. Loading of lipobeads prepared by hydrogel/liposome mixing*

In the case of lipobeads prepared by hydrogel/liposome mixing, hydrophilic cargos usually are introduced into the interior of hydrogel particles, whereas hydrophobic ones—into the lipid bilayer of liposomes before their mixing. Some molecules can penetrate through the lipid bilayer into the hydrogel core.

Back in 1987, the 200–600-nm agarose-gelatin nanogels were filled with colloidal gold particles prior to mixing with liposomes to form lipobeads [2]. Since colloidal gold particles are very adsorptive for proteins and peptides, their encapsulation into hydrogel core can increase the loading capacity of lipobeads. The cytokine Interleukin-2 (IL-2) plays an important role as an immunostimulator and can be relevant as a treatment by itself for cancer and HIV. However, the difficulties faced today with IL-2 are its toxicity and short half-life. To resolve these problems, this protein was bound to the lipobeads (polysaccharide hydrogel nanoparticles coated with lipid bilayer) [4, 93]. In principle, the lipobeads can be loaded with a number of entities just by their incubation in the corresponding solutions, for example, Ca2+ ions and drug mimicking molecules [8], adenosine triphosphate (ATP) [35], and dextran (1.5–3.0 kDa) [44]. Herein, it was found that permeability of the lipid membrane was similar to the free bilayer. To incorporate transmembrane proteins into the peripheral membrane of lipobeads, the hydrogels core particles were mixed and incubated with liposomes containing the proteins of interest within their lipid bilayer [45].

The only chemotherapeutic drug—doxorubicin—was loaded into lipobeads [6, 43]. Encapsulation was performed before lipobeads formation by soaking the dry hydrogel particles in a drug-dissolved solution. The drug diffuses inside in the course of the polymer network swelling and mesh size increase. Further mixing of hydrogel particles with liposomes encapsulates the drug into the lipobeads. As a result, the unbelievably high doxorubicin concentration of ~2 M, which is 10-fold the concentration in liposomes [94], was achieved.

A lecithin-based microemusion method was proposed for fabrication and loading of single-core or multicore lipobeads [31]. The loading efficiency of caffeine into thus prepared sodium hyaluronate lipobeads was obtained to be 30%. A concentration of natural moisturizing factor close to the one present in corneocytes (15%) was encapsulated into the lipobeads, which acquire an enhanced water retention ability similar to corneocytes. This makes them potential for applications in cosmetics and dermatology.

Encapsulation of a protein drug into hydrogel particles before lipobeads formation can be performed either by formation of a hydrogel particle in the presence of a protein drug or by incubation of the preformed hydrogel particles in a protein solution. The first approach again could be problematic due to a danger of protein denaturation. The second approach is limited by the size-exclusion effect resulting in a lower loading concentration of proteins. However, encapsulation of proteins into microgels is a promising tool to increase the amount of drug loaded in a prelipobead (loading capacity) by using the "intelligent" properties of polymer networks (swelling/shrinking ability in response to stimuli) [95].

## **3. Hydrogel core swelling/collapsing and lipobeads properties**

The three-dimensional polymer network within a closed lipid bilayer (liposome) can be considered as a gigantic single molecule stabilized by chemical (covalent bonds) or/and physical (ionic bonds, entanglements, crystallites, charge complexes, hydrogen bonding, van der Waals, or hydrophobic interactions) cross-links. The hydrogel core is also an open container with semipermeable boundaries, across which water and solute molecules can move whereas charged (ionizable) groups fixed on the network chains cannot move (**Figure 4**). Herein, the network of cross-linked polymers exhibits both liquid-like and solid-like behavior [96–99]. Therefore, because of its high water content and elastic nature similar to natural tissue, hydrogel core within a liposome is solid enough to support lipid bilayers and liquid enough to keep the membrane intact and functional.

Besides the mechanical stability that hydrogel core provides to the lipid bilayer, the stimulisensitivity of the polymer network can be used for managing the environmental responsiveness of lipobeads.

#### **3.1. A variety of possible hydrogel cores**

by 10% higher than those for liposomal carriers. A high encapsulation efficiency of lipobeads has been demonstrated also for hemoglobin [14, 15], which withstands the conditions of free radical polymerization and UV radiation. The other example of successful coencapsulation of hydrophilic proteins (IL-2) and small hydrophobic molecules (TGF-receptor-I inhibitor, SB505124) into the biodegradable hydrogel core of lipobeads has been presented in [11]: encapsulation efficiencies were 80 and 36%, respectively, and UV polymerization did not compromise bioactivity of both immunomodulators. Recently, PAAm lipobeads with a good encapsulation efficiency (37%) of enzymes (bovine Cu, Zn-superoxide dismutase, and bovine milk lactoperoxidase) were synthesized to prove that the UV irradiation used for interior gelation did not cause any reduction in the

In the case of lipobeads prepared by hydrogel/liposome mixing, hydrophilic cargos usually are introduced into the interior of hydrogel particles, whereas hydrophobic ones—into the lipid bilayer of liposomes before their mixing. Some molecules can penetrate through the

Back in 1987, the 200–600-nm agarose-gelatin nanogels were filled with colloidal gold particles prior to mixing with liposomes to form lipobeads [2]. Since colloidal gold particles are very adsorptive for proteins and peptides, their encapsulation into hydrogel core can increase the loading capacity of lipobeads. The cytokine Interleukin-2 (IL-2) plays an important role as an immunostimulator and can be relevant as a treatment by itself for cancer and HIV. However, the difficulties faced today with IL-2 are its toxicity and short half-life. To resolve these problems, this protein was bound to the lipobeads (polysaccharide hydrogel nanoparticles coated with lipid bilayer) [4, 93]. In principle, the lipobeads can be loaded with a number of entities just by their incubation in the corresponding solutions, for example, Ca2+ ions and drug mimicking molecules [8], adenosine triphosphate (ATP) [35], and dextran (1.5–3.0 kDa) [44]. Herein, it was found that permeability of the lipid membrane was similar to the free bilayer. To incorporate transmembrane proteins into the peripheral membrane of lipobeads, the hydrogels core particles were mixed and incubated with liposomes containing

The only chemotherapeutic drug—doxorubicin—was loaded into lipobeads [6, 43]. Encapsulation was performed before lipobeads formation by soaking the dry hydrogel particles in a drug-dissolved solution. The drug diffuses inside in the course of the polymer network swelling and mesh size increase. Further mixing of hydrogel particles with liposomes encapsulates the drug into the lipobeads. As a result, the unbelievably high doxorubicin concentration of ~2 M, which

A lecithin-based microemusion method was proposed for fabrication and loading of single-core or multicore lipobeads [31]. The loading efficiency of caffeine into thus prepared sodium hyaluronate lipobeads was obtained to be 30%. A concentration of natural moisturizing factor close to the one present in corneocytes (15%) was encapsulated into the lipobeads, which acquire an enhanced water retention ability similar to corneocytes. This makes them potential for applica-

enzymatic activity of the proteins [37].

62 Liposomes

lipid bilayer into the hydrogel core.

*2.5.2. Loading of lipobeads prepared by hydrogel/liposome mixing*

the proteins of interest within their lipid bilayer [45].

is 10-fold the concentration in liposomes [94], was achieved.

tions in cosmetics and dermatology.

Depending on the composition of a gel/solvent system, the polymer and cross-linking chemistry, nanogels swell or shrink discontinuously or continuously, reversibly or irreversibly in response to many different stimuli (temperature, pH, ion concentration, electric fields, light, reduction/oxidation, enzymatic activity, etc.) [47, 100–105]. In general, various types of hydrogels based upon either natural (e.g., hyaluronic acid, collagen, chondroitin sulfate, alginates, fibrin, and chitosan) and synthetic polymers made of neutral (e.g., 2-hydroxyethyl methacrylate, N-alkylmethacrylamides, N-alkylacylamides, and N,N-dialkylacrylamides), acidic (e.g., acrylic acid, metacrylic acid, and 2-acrylamido-2-methyl propane sulfonic acid), or basic (e.g., N,N-dialkylaminoethyl methacrylate, 1-vinylimidasole, and methacryloyoloxyethyltrialkylammonium bromide) monomers have been prepared, studied, and used in numerous applications (bioseparation, tissue engineering, sensing and molecular recognition, drug and gene delivery, controlled release, artificial muscles, and flow control).

#### *3.1.1. Temperature-sensitive volume change*

Typically, the thermoresponsive hydrogels are classified as having either positive or negative volume phase transition with a characteristic temperature (*T*V) [106]. Hydrogels exhibiting positive volume phase transition ("thermophilic" hydrogels) swell upon heating. In contrast, hydrogels exhibiting negative volume-phase transition ("thermophobic" hydrogels) collapse upon heating. The "thermophobic" hydrogels have been studied the most, and a popular example is poly(*N*-isopropylacrylamide) (PNIPA) [107]. In contrast, "thermophilic" behavior in water is not very common for synthetic polymeric materials and likely, because of that, lipobeads with a "thermophilic" hydrogel core have not been attempted yet. Nevertheless, a number of "thermophilic" hydrogels showing a positive thermosensitive volume change have been already fabricated [108–110]. Chitosan cross-linked with glutaraldehyde was found to exhibit the swelling behavior in aqueous media at physiological temperatures and pHs [111]. The graft copolymerization of mixtures of acrylamide (AAm) and acrylonitrile (AN) with *Gum ghatti* (Gg) and cross-linking with MBA resulted in a hydrogel capable of twofold swelling when temperature raised from 30 to 40°C in distilled water [112]. The interpenetrating polymer networks (IPN) composed of poly(acrylic acid) (PAAc) and poly(acrylamide(AAm)-co-butyl methacrylate (BMA), as well as the random poly(AAm-co-AAc-co-BMA) hydrogels cross-linked with MBA also increased their volume (~2.5-fold) within the 30–40°C range in water [113]. Positive thermosensitivity with a twofold swelling ability in the range from 30 to 45°C was reported for a nonionic chemically cross-linked gel made of *N*-acryloylglycinamide as a monomer and MBA as a cross-linker [109]. An abrupt increase in volume to the similar extent within the interval of 30–40°C was observed in the polyzwitterionic hydrogels consisting of N,N′ dimethyl(methacroylethyl)ammonium propanesulfonate or N,N′-dimethyl(acrylamidopropyl) ammonium propanesulfonate cross-linked with EGDM [114]. The gels with positive volume transitions at physiological temperatures, pH and in the presence of salt can be useful in biological or biomedical applications. However, in all abovementioned cases, under physiological pH and salt concentrations, the hydrogels either do not show thermophilic behavior at all or the transition temperature shifted to the nonphysiological values. Nonetheless, recently, hydrogels based on the cross-linked poly(allylurea-co-allylamine) (PAU) copolymers were prepared to unveil a fast and pronounced "thermophilic" increase in volume within the physiological ranges of temperature, pH, and concentration of salt [110].

Thus, first and foremost, a temperature range where the hydrogel shrinks or swells intrinsically depends on the chemical nature of the polymer constituting its network. Herein, the volume changes in a water-swollen hydrogel can be either continuous or discontinuous, as a function of environmental stimuli. If the system remains totally miscible at given thermodynamic conditions, one can expect continuous volume transition. On the contrary, if changes in chemical nature of the polymer network, solvent quality, or environmental stimulus "push" the system into a two-phase (unstable) region of the solubility phase diagram, one can expect that properties of the hydrogel, most notably its volume, change discontinuously. In addition, the studies [115–117] show that increased cross-linking may significantly decrease swelling ability of hydrogel, especially, below *T*V, but has a little effect on the value of *T*V.

**Figure 6** demonstrates that temperature-sensitive shrinking ability of hydrogels depends on their microstructure and method of preparation: the granular hydrogel (A) exhibits a continuous volume decrease with temperature, whereas the denser hydrogels (B and C) exhibit more abrupt changes in volume within the 35–45°C range. Interestingly, the granular structure of the PNIPA hydrogels prepared by thermal polymerization in water (A) can be broken down into separate submicroscopic domains by means of sonication (data not shown). On

positive volume phase transition ("thermophilic" hydrogels) swell upon heating. In contrast, hydrogels exhibiting negative volume-phase transition ("thermophobic" hydrogels) collapse upon heating. The "thermophobic" hydrogels have been studied the most, and a popular example is poly(*N*-isopropylacrylamide) (PNIPA) [107]. In contrast, "thermophilic" behavior in water is not very common for synthetic polymeric materials and likely, because of that, lipobeads with a "thermophilic" hydrogel core have not been attempted yet. Nevertheless, a number of "thermophilic" hydrogels showing a positive thermosensitive volume change have been already fabricated [108–110]. Chitosan cross-linked with glutaraldehyde was found to exhibit the swelling behavior in aqueous media at physiological temperatures and pHs [111]. The graft copolymerization of mixtures of acrylamide (AAm) and acrylonitrile (AN) with *Gum ghatti* (Gg) and cross-linking with MBA resulted in a hydrogel capable of twofold swelling when temperature raised from 30 to 40°C in distilled water [112]. The interpenetrating polymer networks (IPN) composed of poly(acrylic acid) (PAAc) and poly(acrylamide(AAm)-co-butyl methacrylate (BMA), as well as the random poly(AAm-co-AAc-co-BMA) hydrogels cross-linked with MBA also increased their volume (~2.5-fold) within the 30–40°C range in water [113]. Positive thermosensitivity with a twofold swelling ability in the range from 30 to 45°C was reported for a nonionic chemically cross-linked gel made of *N*-acryloylglycinamide as a monomer and MBA as a cross-linker [109]. An abrupt increase in volume to the similar extent within the interval of 30–40°C was observed in the polyzwitterionic hydrogels consisting of N,N′ dimethyl(methacroylethyl)ammonium propanesulfonate or N,N′-dimethyl(acrylamidopropyl) ammonium propanesulfonate cross-linked with EGDM [114]. The gels with positive volume transitions at physiological temperatures, pH and in the presence of salt can be useful in biological or biomedical applications. However, in all abovementioned cases, under physiological pH and salt concentrations, the hydrogels either do not show thermophilic behavior at all or the transition temperature shifted to the nonphysiological values. Nonetheless, recently, hydrogels based on the cross-linked poly(allylurea-co-allylamine) (PAU) copolymers were prepared to unveil a fast and pronounced "thermophilic" increase in volume within the physiological

Thus, first and foremost, a temperature range where the hydrogel shrinks or swells intrinsically depends on the chemical nature of the polymer constituting its network. Herein, the volume changes in a water-swollen hydrogel can be either continuous or discontinuous, as a function of environmental stimuli. If the system remains totally miscible at given thermodynamic conditions, one can expect continuous volume transition. On the contrary, if changes in chemical nature of the polymer network, solvent quality, or environmental stimulus "push" the system into a two-phase (unstable) region of the solubility phase diagram, one can expect that properties of the hydrogel, most notably its volume, change discontinuously. In addition, the studies [115–117] show that increased cross-linking may significantly decrease swelling

**Figure 6** demonstrates that temperature-sensitive shrinking ability of hydrogels depends on their microstructure and method of preparation: the granular hydrogel (A) exhibits a continuous volume decrease with temperature, whereas the denser hydrogels (B and C) exhibit more abrupt changes in volume within the 35–45°C range. Interestingly, the granular structure of the PNIPA hydrogels prepared by thermal polymerization in water (A) can be broken down into separate submicroscopic domains by means of sonication (data not shown). On

ability of hydrogel, especially, below *T*V, but has a little effect on the value of *T*V.

ranges of temperature, pH, and concentration of salt [110].

64 Liposomes

**Figure 6.** Shrinking abilities *S*V and SEM images of the microstructures of PNIPA hydrogels prepared by thermal polymerization in water (A) or DMSO (B), and by UV polymerization in water (C) (scale bar = 5 μm).

the contrary, sonication does not affect the structure of "dense" hydrogels prepared either by thermal polymerization in dimethyl sulfoxide (DMSO) (B) or by photopolymerization in water (C).

It has been shown [118] that incorporation of a small amount of ionizable groups into the nonionic gel network drives the volume phase transition from continuous volume changes toward discontinuous one (**Figure 7**). Moreover, an increase in the portion of sodium acrylate with carboxylic groups on the PNIPA network allowed one to vary the *T*V from 34 to 42°C with the increasing extent of swelling ability below the transition temperature.

**Figure 7.** The degree of swelling of the poly(*N*-isopropylacrylamade-co-sodium acrylate) gel in water as a function of temperature. Numbers are the molar fractions of sodium acrylate in the preparations. Data were adopted from [118].

#### *3.1.2. Ionic sensitivity of hydrogels*

**Figure 7** also suggests that incorporation of charged (anionic, cationic, or both) groups on the polymer network makes the volume transition temperature and degree of swelling dependent on pH and ionic strength. Indeed, the poly(N-isopropylacrylamide-co-methacrylic acid) (PNIPA-MAA) microgel particles [119] at pH 3.4 exhibited a decrease in *T*V from 33.5 to 28°C with an increase in MAA content, whereas at pH 7.5, the higher MAA content resulted in the higher *T*V. In weakly charged PNIPA hydrogels, addition of ionizable groups on the polymer network pronounced the volume changes when temperature crossed *T*V [118, 120, 121]. The experimental studies [122] revealed that distribution of ionic groups in the network affects the temperature of volume change transition.

The type of ionizable groups on the polymer networks makes the maximum swelling ability of a gel strongly dependent on pH. For example, the anioic PNIPA-MAA microgels exhibited the maximum swelling in the range of pHs from 6.5 to 10 [119], whereas for the cationic PNIPA-VI nanogels, the maximum swelling ratio was observed in the range of pHs from 6 to 3.5 [123]. It becomes even more intriguing if the so-called polyampholyte hydrogels are designed [124] by addition of both cationic (VI) and anionic (AA) groups on the network. The polyampholyte gel was in a shrunken state near the isoelectric point (pH ~ pI), and it swelled at both higher and lower pHs. It is interesting that such designed polyampholyte gels can work like biochemomechanical systems in which the enzymatically induced pH changes control the volume of polyampholyte network or, in opposite direction, the pH sensitive volume changes control the activity of enzymes immobilized into the gel [125].

There are experimental evidences [126–130] that different monovalent (Li+ , Na+ , K+ , and Cs+ ) and divalent (Ca2+, Mg2+, Sr2+, and Ba2+) ions are able to promote deswelling effects in hydrogels of different chemical nature. Interestingly, at the same molar ratios of divalent to monovalent cations (~1 mM/30 mM), the similar volume changes were observed in biological polyelectrolyte systems during physiological processes like nerve excitation, muscle contraction, and cell locomotion [131–137].

#### *3.1.3. Surfactants as effectors of hydrogel volume change*

The extensive theoretical and experimental [138–145] studies have shown that the addition of anionic, cationic, and nonionic surfactants to the solution containing a gel can also influence the *T*V and swelling degree of hydrogels depending on their hydrophobicity and charge of the polymer network. In general, addition of anionic or cationic surfactant to the solution of nonionic hydrogel increases *T*V as well as the swelling range, whereas the nonionic surfactant does not affect *T*V or volume change. The surfactants with ionic head groups when bind to the nonionic polymer networks convert the neutral hydrogels to polyelectrolyte gels and elevate *T*V due to introduction of additional osmotic pressure by ionization. The changes in the volume phase transition are also dependent on the length of hydrophobic tail of ionic surfactants and the critical concentration of micelle formation [142]. It was also found [144] that the amount of an ionic surfactant bound onto the swollen network of the nonionic PNIPA hydrogel was much greater than that to the collapsed one. On the contrary, the amount of nonionic surfactant bound onto the collapsed network of the PNIPA gel was greater than that on the swollen one. Moreover, the changes in *T*V with the amount of the anionic surfactants (e.g., sodium dodecyl sulfonate) were more pronounced than for cationic ones (e.g., dodecylamine hydrochloride). Interestingly, the concentration of anionic surfactant (e.g., sodium dodecylbenzene sulfate) bound within the PNIPA hydrogel was found to be higher in the vicinity of the gel surface, whereas a central region of the gel may not contain any bound surfactant molecules [145]. Thus, peripheral layers could be in a more swollen state with a higher *T*V in comparison to the central hydrogel core.

#### *3.1.4. Light-sensitive hydrogels*

*3.1.2. Ionic sensitivity of hydrogels*

66 Liposomes

volume change transition.

locomotion [131–137].

activity of enzymes immobilized into the gel [125].

*3.1.3. Surfactants as effectors of hydrogel volume change*

There are experimental evidences [126–130] that different monovalent (Li+

**Figure 7** also suggests that incorporation of charged (anionic, cationic, or both) groups on the polymer network makes the volume transition temperature and degree of swelling dependent on pH and ionic strength. Indeed, the poly(N-isopropylacrylamide-co-methacrylic acid) (PNIPA-MAA) microgel particles [119] at pH 3.4 exhibited a decrease in *T*V from 33.5 to 28°C with an increase in MAA content, whereas at pH 7.5, the higher MAA content resulted in the higher *T*V. In weakly charged PNIPA hydrogels, addition of ionizable groups on the polymer network pronounced the volume changes when temperature crossed *T*V [118, 120, 121]. The experimental studies [122] revealed that distribution of ionic groups in the network affects the temperature of

The type of ionizable groups on the polymer networks makes the maximum swelling ability of a gel strongly dependent on pH. For example, the anioic PNIPA-MAA microgels exhibited the maximum swelling in the range of pHs from 6.5 to 10 [119], whereas for the cationic PNIPA-VI nanogels, the maximum swelling ratio was observed in the range of pHs from 6 to 3.5 [123]. It becomes even more intriguing if the so-called polyampholyte hydrogels are designed [124] by addition of both cationic (VI) and anionic (AA) groups on the network. The polyampholyte gel was in a shrunken state near the isoelectric point (pH ~ pI), and it swelled at both higher and lower pHs. It is interesting that such designed polyampholyte gels can work like biochemomechanical systems in which the enzymatically induced pH changes control the volume of polyampholyte network or, in opposite direction, the pH sensitive volume changes control the

and divalent (Ca2+, Mg2+, Sr2+, and Ba2+) ions are able to promote deswelling effects in hydrogels of different chemical nature. Interestingly, at the same molar ratios of divalent to monovalent cations (~1 mM/30 mM), the similar volume changes were observed in biological polyelectrolyte systems during physiological processes like nerve excitation, muscle contraction, and cell

The extensive theoretical and experimental [138–145] studies have shown that the addition of anionic, cationic, and nonionic surfactants to the solution containing a gel can also influence the *T*V and swelling degree of hydrogels depending on their hydrophobicity and charge of the polymer network. In general, addition of anionic or cationic surfactant to the solution of nonionic hydrogel increases *T*V as well as the swelling range, whereas the nonionic surfactant does not affect *T*V or volume change. The surfactants with ionic head groups when bind to the nonionic polymer networks convert the neutral hydrogels to polyelectrolyte gels and elevate *T*V due to introduction of additional osmotic pressure by ionization. The changes in the volume phase transition are also dependent on the length of hydrophobic tail of ionic surfactants and the critical concentration of micelle formation [142]. It was also found [144] that the amount of an ionic surfactant bound onto the swollen network of the nonionic PNIPA hydrogel was much greater than that to the collapsed one. On the contrary, the amount of nonionic

, Na+ , K+

, and Cs+

)

Photosensitive hydrogels with incorporated photosensitive molecules into the gel network have been reported as well. For example, the gels with incorporated leucocyanide and leucohydroxide [146] underwent volume changes upon irradiation and removal of ultraviolet light resulted from ionization reaction and internal osmotic pressure initiated by UV light. Significant volume changes in hydrogels were also induced by visible light [147]. However, the mechanism of volume transition was different—it was due to direct heating of the polymer network by light. Nevertheless, more recent reports [148] showed that a focused laser beam was able to induce reversible shrinking in polymer gels due to radiation forces, rather than local heating, modifying the weak interactions in the gels. Herein, gel shrinkage was observed up to several tens of micrometers away from the irradiation spot. The lightinduced contraction was also found in acrylamide gels, which are not temperature sensitive. In hydrogels with temperature-sensitive volume phase transitions, such as PNIPA gel, it was found that the radiation force of the laser beam not only induces the volume phase transition but also lowers the transition temperature *T*V by about 10°C at an irradiation power of 1.2 W (*λ* = 1064 nm).

The fact that the volume change initiated by light is extremely fast seems of great importance for the development of the light-sensitive lipobeads. One could predict that the photosensitive hydrogel cores will also receive an increasing scientific and technological attention due to their capability of serving as the so-called shape-memory polymeric systems [149]. Being exposed to the light with lower wavelengths, the shape-memory materials become deformed and their temporary shape is fixed due to cross-linking. When irradiated with higher wavelengths, they recover their initial shape because of the cross-links cleavage.

#### *3.1.5. Electrical field-induced volume change*

Back in the 1950s, it was found that contraction, oscillation, and bending of polyelectrolyte gels can be induced electrically [150–152]. In particular, gels prepared from polymers and copolymers that contain ionizable groups exhibited remarkable contraction when placed between a pair of electrodes connected to a direct current source. Polymer gels containing no ionizable groups showed no volume change under electrical field applied. The extent and rate of volume change of the polyelectrolyte gels were shown to increase with increasing electrical field [153]. An increase in the ionic strength (e.g., an addition of NaCl) also increases the rate of gel shrinkage, whereas an addition of organic solvent (e.g., acetone or ethanol) decreases both the extent and the rate of shrinking. In different types of hydrogels, an electrically activated volume changes were associated with the induction of the medium pH change by the electric field [154], the electrically initiated volume phase transition [155], and the so-called electrokinetic phenomena—ion transport of counter-ions in the electric field [156].

#### **3.2. Lipobeads by hydrogel/liposome mixing**

Success in formation of lipobeads by hydrogel/liposome mixing (see references from **Tables 3** and **4**) is an experimental confirmation of the main property of hydrogel and lipid bilayer their compatibility. Indeed, the phospholipid bilayer spontaneously self-assembled around a nanogel once extracted from a lipobead (**Figure 8a**) and mixed with liposomes (**Figure 8b**) to form a secondary lipobead (**Figure 8c**) [10].

Moreover, spontaneous formation of the lipid bilayer on the surface of nonanchored microgels was shown microscopically for liposomes made of different phospholipids with or without cholesterol [36, 41]. **Figure 9** represents both laser scanning confocal (**Figure 9A**, **A′**, **A"**) and scanning electron microscopy (**Figure 9B**, **B´**) images evidencing the fusion of liposomes on the hydrogel surface to form a lipid membrane around porous PNIPA-co-FA microgels.

Unilamellarity, continuity, and nonleakiness of the lipid bilayer formed upon microgel/liposome mixing were proven for the micrometer-sized, hydrophobically modified, pNIPAM/p(NIPAMco-AA) core-shell hydrogel spheres [25, 28]. It was also demonstrated by Dynamic Light Scattering (DLS) and Atomic Force Microscopy (AFM) [40] that hydrophobic modification of the nanogels is not required for spontaneous formation of the bilayer on their surface.

Interesting behavior of the lipobeads was observed when their hydrogel cores change its volume. It was shown (**Figure 10**) that a collapse of the hydrophobically modified PNIPA microgels within the lipid bilayer caused a shape change of the lipobeads from sphere (below *T*V) to a small

**Figure 8.** Atomic force microscopy images (amplitude data) of a PNIPA-VI nanogel (a), EPC liposomes (b), and result of their mixture—lipobead flattened on mica surface. Scale bars = 100 nm.

ethanol) decreases both the extent and the rate of shrinking. In different types of hydrogels, an electrically activated volume changes were associated with the induction of the medium pH change by the electric field [154], the electrically initiated volume phase transition [155], and the so-called electrokinetic phenomena—ion transport of counter-ions in the electric

Success in formation of lipobeads by hydrogel/liposome mixing (see references from **Tables 3** and **4**) is an experimental confirmation of the main property of hydrogel and lipid bilayer their compatibility. Indeed, the phospholipid bilayer spontaneously self-assembled around a nanogel once extracted from a lipobead (**Figure 8a**) and mixed with liposomes (**Figure 8b**) to

Moreover, spontaneous formation of the lipid bilayer on the surface of nonanchored microgels was shown microscopically for liposomes made of different phospholipids with or without cholesterol [36, 41]. **Figure 9** represents both laser scanning confocal (**Figure 9A**, **A′**, **A"**) and scanning electron microscopy (**Figure 9B**, **B´**) images evidencing the fusion of liposomes on the hydrogel surface to form a lipid membrane around porous PNIPA-co-FA

Unilamellarity, continuity, and nonleakiness of the lipid bilayer formed upon microgel/liposome mixing were proven for the micrometer-sized, hydrophobically modified, pNIPAM/p(NIPAMco-AA) core-shell hydrogel spheres [25, 28]. It was also demonstrated by Dynamic Light Scattering (DLS) and Atomic Force Microscopy (AFM) [40] that hydrophobic modification of

Interesting behavior of the lipobeads was observed when their hydrogel cores change its volume. It was shown (**Figure 10**) that a collapse of the hydrophobically modified PNIPA microgels within the lipid bilayer caused a shape change of the lipobeads from sphere (below *T*V) to a small

**Figure 8.** Atomic force microscopy images (amplitude data) of a PNIPA-VI nanogel (a), EPC liposomes (b), and result of

their mixture—lipobead flattened on mica surface. Scale bars = 100 nm.

the nanogels is not required for spontaneous formation of the bilayer on their surface.

field [156].

68 Liposomes

microgels.

**3.2. Lipobeads by hydrogel/liposome mixing**

form a secondary lipobead (**Figure 8c**) [10].

**Figure 9.** The bright field (A) and confocal laser scanning (A′, A″) microscopy images of PNIPA-co-FA hydrogel microspheres mixed with the liposomes made of HSPC with cholesterol (molar ratio 9:1): Green image (A′) originated from fluorescein-o-acrylate (FA) covalently attached to the PNIPA network within the core. Red image (A″) originated from rhodamine B covalently attached to the heads of PE. Scanning electron micrographs of the PNIPA-co-FA microgel before (B) and after (B′) mixing with liposomes (scale bars = 20 μm).

**Figure 10.** Fluorescence images of a giant anchored PNIPA lipobead below (left) and above (right) Tv. Red images originated from 0.05 mol% RhodB-PE component within the lipid bilayer. The images were adopted from [25, 28].

central core with high curvature protrusions (above *T*V), consisting of the excess lipid bilayer which still adjoin the lipid bilayer remaining bound to the hydrogel via hydrophobic anchors [25, 28]. Importantly, these changes were found to be reversible, and the bilayer remained intact and impermeable.

#### **3.3. Lipobeads by polymerization within liposomal interior**

#### *3.3.1. Lipobeads with hydrophobically modified nanogels*

As shown by DLS, the size distribution of lipobeads with hydrophobically anchored PNIPA-VI nanogels became bimodal when temperature was raised to 40°C: the position of the first peak corresponded to the initial size of lipobeads at 25°C, while the second peak was assigned to the aggregates of lipobeads. The further cooling back to 25°C restored the original unimodal size distribution of lipobeads, indicating reversibility of anchored lipobeads aggregation. **Figure 11** sketches the behavior of lipobeads resulted from the anchored hydrogel collapsing.

Presumably, the aggregation reduces the hydrophobic/hydrophilic imbalance caused by collapsed nanogels within lipobeads. The reversible dissociation of the lipobead aggregates may evidence that anchored lipobeads do not fuse. The only explanation is that the hydrophobic chains of anchored PNIPA-VI nanogels penetrate into the lipid bilayer and stabilize the liposomal membrane against fusion.

On the contrary to the hydrophobically modified giant lipobeads (**Figure 10**), the anchored PNIPA-VI nanolipobeads did not reveal a size change under temperature or pH variations, as shown by DLS [9]. Probably, on the nanometer scale, a highly curved lipid bilayer is too stiff to follow the collapse of the hydrogel core. This effect remains to be proven.

#### *3.3.2. Lipobeads with nonmodified nanogels*