Emergence of Aedes aegypti adults with eggs incubated for 10 days at different temperatures and placed for post-embryonic development at a temperature of 25 ± 2°C.
\n\n
\n\nThe research leading to these results has received funding from the European Community\'s Seventh Framework Programme (FP7/2007-2013) under grant agreement number 285417. The publishing of this book was funded by the EC FP7 Post-Grant Open Access Pilot programme. 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In addition, the severe dengue is one of the main causes of child deaths in countries in Latin America and Asia [3]. The circulation of the dengue virus is ruled as a major problem due to the genetic plasticity of the mosquito under environmental conditions, and due to the selection of resistant insects because of chemical control, coupled with the fact that the mosquito is anthropophilic and domiciled. The occurrence of vertical transmission in epidemic times also helps in the dynamics of maintaining the virus in the urban environment [4]. Murillo et al. [5] reported that increased vertical transmission in times of epidemic makes outbreaks more difficult and expensive to control.
Aedes (Stegomyia) aegypti (Diptera: Culicidae) is the transmitter of dengue, urban yellow fever, chikungunya and Zika viruses in Brazil [6, 7]. The spread of chikungunya and Zika viruses has caused public health problems because the former causes severe recurrent joint pain and the latter may involve neurological complications such as microcephaly in children, being very severe, in addition to Guillain-Barré Syndrome [8, 9, 10, 11].
Females of this mosquito species practice hematophagy and prefer to breed in artificial breeding sites, where competition with other species is practically absent [12, 13, 14, 15]. The proliferation of this mosquito has been intensifying with increasing urban perimeter, lack of basic sanitation, deforestation and lack of proper treatment of solid waste, as these are factors that increase breeding sites [16, 17]. A. aegypti eggs can be dehydrated for more than a year and are viable in late-hatching post-embryonic development, causing an imminent danger in the proliferation of this mosquito [18, 19].
Mosquito development, especially those with high adaptive genetic plasticity, may be affected by various abiotic factors, with temperature being the most relevant [20]. According to the Intergovernmental Panel on Climate Change [21], the Earth will experience an average temperature increase of approximately 2°C, and by 2099 this could rise to 6.5°C, a situation that may favor the cycle of these Culicidae, as well as increased viral circulation. These data are confirmed by analyzing the statistics from 1955 to 2007, a time when temperature increases have been added to one of the world’s largest viral propagations, with a 30-fold increase in the last 50 years, and over 2 million of reported cases annually on the American continent each year [3].
In the current perspective, there is a need to intensify the monitoring and control of mosquitoes, as well as the monitoring of early viral circulation, since almost half of the world’s population is exposed to the risk of dengue contamination with an estimated 50–100 million cases annually in more than 100 endemic countries [3]. This situation can worsen with global warming, which has been increasing each year, due to anthropic factors [17]. In this sense, understanding the life cycle of A. aegypti in stabilized insectaries kept in environmental rooms simulating the IPCC climate predictions, is a strategy that can help to optimize the alternatives for monitoring and controlling of this insect, given the possibility of the emergence of this scenario.
In this context, the present work aimed to: (i) verify the influence of different temperatures on egg viability, number of adults and mortality rate of A. aegypti from Londrina, Paraná, under laboratory conditions and, (ii) check the time of the biological cycle, number of adults and mortality rate of A. aegypti from Manaus, Amazonas, kept in environmental rooms that simulate the temperatures and concentrations of carbon dioxide predicted by the IPCC.
The study was carried out in incubator chambers (BOD) with different temperatures, kept at the Medical Entomology Laboratory of Londrina State University (UEL), Londrina, Paraná, and in insects kept at the Laboratory of Ecophysiology and Molecular Evolution (LEEM), located at Campus I of the National Institute of Amazonian Research (INPA), Manaus, Amazonas.
The sample of A. aegypti population from Londrina, Paraná, used in this study, was obtained from eggs collected in the field, with the aid of traps called ovitraps [22]. The main attraction in the traps was a solution containing grass infusion (10%) [23], with a total volume of 300 ml. Twenty traps were set up and distributed at the UEL campus, at ground level, protected from sun and rain, and in places with little movement of people and animals. Samples were taken for 2 weeks and every 7 days, the reeds were replaced and sent to the insectarium with controlled temperature, humidity and photoperiod conditions (27 ± 2°C, 80–90% and 12L/12E).
The reeds containing eggs were immersed in plastic trays (45 × 30 × 7.5 cm) containing distilled water to stimulate the larvae to hatch. The immatures obtained were kept until adults by means of food containing a mixture of cat food (Whiskas®) and rodents (Teklab global®) in a 1:1 ratio, ground into fine particles (1 mm). All trays were covered with a nylon fabric to prevent the escape of mosquitoes. After emergence, the males and females were collected with a Castro catcher and the species identification was performed using external morphological characters, mainly from the chest, with the aid of a stereoscopic microscope ZEISS Stemi 2000 50× and identification keys proposed by Forattini [14], Harbach [24], and WRBU [25].
After the identification stage, the adults were placed to copulate in cardboard cages (17 × 20 cm), containing two plastic cups lined with strips of filter paper and filled with 70 mL of distilled water, which were used as a substrate for oviposition. As a source of carbohydrates, an Erlenmeyer was introduced containing a roll of gauze with pieces of cotton in the center, soaked with 12% sugar water. The blood meal was carried out using an anesthetized hamster for 30min, according to the procedure approved by the Ethics Committee on the Use of Animals at UEL (“Breeding of mosquitoes in laboratory conditions”).
At UEL, the experiment was carried out in four incubator chambers (BOD) that had different temperatures (0, 5, 25 and 45°C) with ±2°C, as well as at ambient temperature, where the eggs remained in dry incubation in 500 ml capacity pots containing only moistened cotton over a period of 10 days, with a 12/12 h photoperiod.
Subsequently, 150 ml of distilled water, 25 eggs of A. aegypti, F1 field generation and 0.055 g of larval food were introduced into each of the five pots, which were placed in BOD with temperature of 25 ± 2°C. The monitoring of the hatching rate of eggs, the rate of immature deaths and the number of adults were carried out daily.
The same process of creating and maintaining the immature and adults of A. aegypti from Londrina, Paraná, mentioned in items 2.2 and 2.3, was carried out with a sample of the population of A. aegypti from Manaus, Amazonas, collected in the same way on Campus I of INPA. However, the blood meal carried out followed the procedure approved by the Ethics Committee on the Use of Animals at INPA (CEUA/INPA: 04/2013-“Breeding of mosquito vectors, under laboratory conditions”).
After the stabilization of the three insectaries in the LEEM, the F28 generation was obtained in the different environmental rooms that started the experiment for post-embryonic development, using three different environments. Rooms 1, 2 and 4 suffer from interference from different temperatures and levels of carbon dioxide, and rooms 2 and 4 refer to 50 and 100 years in front of room 1, which presents ambient temperature.
These rooms are equipped with technological devices that guarantee, respectively, the maintenance of ambient temperature, 2.5°C (400 ppm CO2) and 4.5°C (850 ppm CO2) above ambient temperature. The natural conditions of the respective environmental rooms are collected in real time by sensors isolated in the forest and the humidity of all remains constant in relation to the environmental conditions.
After stabilization of insectaries in environmental rooms 1 (ambient temperature), 2 (mild temperature) and 4 (extreme temperature), 125 eggs from each room were counted with the aid of a stereoscopic microscope. These eggs were separated into 5 groups (replicates) of 25 eggs, kept in pots with capacity of 750 mL, containing 300 mL of distilled water and 0.055 g of larval food added every 3 days.
The hatching rate was monitored daily for 19 days under these conditions. The average time of the biological cycle, number of adults (males and females) and larval mortality rate were verified. Exuvia and dead immatures were removed from the pots and the volume of distilled water was replaced.
Data on the hatching rate of eggs, number of adults (males and females) and mortality rate of immature A. aegypti from Londrina, Paraná, were expressed as a percentage. The number of emerged adults was subjected to a Kruskal-Wallis test, at the level of 5% significance, in order to verify differences between the data in the temperatures analyzed, using the BioEstat® 5.3 program for Windows [26].
The results on the number of adults (males and females) and the larval mortality rate of A. aegypti from Manaus, Amazonas, were also analyzed by means of percentage. On the other hand, the data regarding the average time of the biological cycle of mosquitoes kept in the different environmental rooms, were analyzed using averages, standard deviation and standard error. At first, these data underwent a Lilliefors normality test (K samples) to find out whether they have a normal distribution or not, and subsequently subjected to Analysis of Variance (ANOVA), followed by Tukey’s multiple comparison test (p ≤ 0.05), with the aid of the statistical program SPSS® 14.0 packpage for Windows® (SPSS Inc. 2005 Headquarters, Chicago, Illinois, USA).
The highest hatching rates of eggs were obtained at temperatures of 5 and 25 ± 2°C, respectively, where 48% for both was verified, followed by ambient temperature with 36% and at 0 ± 2°C presenting 7%. On the other hand, there was no hatching of eggs at 45 ± 2°C (Figure 1).
Egg hatch percentage of A. aegypti from Londrina, Paraná, in incubator chambers (BOD) with different temperatures with ±2°C limits and ambient temperature (16.7–24.1°C), observed for 10 days.
The adults obtained in the proportion of males and females are shown in Table 1.
Adult emergence rate | Temperature (±2°C) | |||||||||
---|---|---|---|---|---|---|---|---|---|---|
0°C | 5°C | 25°C | 45°C | Ambient (16.7–24.1°C) | ||||||
Replicas | M | F | M | F | M | F | M | F | M | F |
1 | 4 | 1 | 8 | 7 | 10 | 4 | 0 | 0 | 6 | 5 |
2 | 0 | 1 | 5 | 4 | 8 | 6 | 0 | 0 | 0 | 0 |
3 | 0 | 0 | 8 | 4 | 5 | 2 | 0 | 0 | 0 | 0 |
4 | 0 | 1 | 9 | 7 | 12 | 3 | 0 | 0 | 10 | 6 |
5 | 0 | 0 | 4 | 0 | 0 | 0 | 0 | 0 | 9 | 8 |
Total | 4 | 3 | 34 | 22 | 35 | 15 | 0 | 0 | 25 | 19 |
Total M-F* | 7a+ | 56a | 50a | 0b | 44a | |||||
Total % | 5.60% | 44.80% | 40% | 0% | 35.20% |
Emergence of Aedes aegypti adults with eggs incubated for 10 days at different temperatures and placed for post-embryonic development at a temperature of 25 ± 2°C.
M-F: M = male and F = female.
Equal letters on the same line do not differ from each other by the Kruskal-Wallis p = 0.0261, H = 11.0428.
At 0 ± 2°C, seven of nine immatures reached adulthood, which indicates a mortality of 22.2% of the larvae at this temperature. Considering the 60 immatures obtained at a temperature of 5 ± 2°C, 56 reached the adult stage indicating a larval mortality of 6.6%, while at a temperature of 25 ± 2°C only 50 immatures among 60 reached the adult stage, indicating a larval mortality of 16.6%. At ambient temperature, it was observed that of 45 immatures obtained, 44 reached adulthood, representing a larval mortality of only 2.2%. On the other hand, there was no adult emergency at 45 ± 2°C.
There is no statistical difference between hatching rates at 0, 5, 25 and ambient temperatures (Table 1), while at 0°C there is difference in relation to the hatching rate at other temperatures.
Of the total 625 eggs incubated at different temperatures, 157 eggs reached adulthood, and of these, 98 were males and 59 females, making up a male:female ratio of 1.66:1. The proportion of emerged adults was 5.6% at 0°C, 44.8% at 5°C, 40% at 25°C and 35.2% at room temperature (Table 1).
The averages of climatic variations during the experiment in °C, increase in ppm of CO2 and percentage (%) of humidity for the different environmental rooms are represented in Table 2.
Rooms | Climate variations | ||
---|---|---|---|
Temperature (°C) | CO2 (ppm) | Humidity (%) | |
1 | 27.23 | 434.4 | 77.01 |
2 | 29.74 | 864.6 | 78.66 |
4 | 32.18 | 1279.87 | 76.96 |
Averages of temperature °C, increase in ppm of CO2 and percentage (%) of humidity for the different environmental rooms.
There was a statistical difference (p < 0.05) between the average time of the biological cycle of A. aegypti in the different environments, with room 4 being the environment that had the greatest influence on the life cycle of mosquitoes, since they developed more faster than the insects kept in the other environmental rooms (Table 3).
Parâmeters | Room 1 (TA) | Room 2 (TM) | Room 4 (TE) |
---|---|---|---|
Averages | 7.3430a,* | 6.6780b,* | 6.0510c,* |
Standard deviation | 0.43756 | 0.65009 | 0.60656 |
Standard error | 0.13837 | 0.20558 | 0.19181 |
Average life cycle time of adults kept in different environmental rooms that simulate the climatic conditions provided by the IPCC.
TA = ambient temperature; TS = mild temperature; TE = extreme temperature. For the Tukey test, the averages were transformed into a root of x + 0.5*. Different letters on the same line, indicate a statistically significant difference according to the Tukey test (p < 0.05), CV = 4%; F = 12.64 and DMS = 0.120.
In the room 1, it was found that adults emerged on the sixth and seventh day, with 45.79% females and 54.20% males. In the room 2, adults emerged on the sixth day, with 58.88% of females and 41.11% of males. In the room 4, adults began to emerge on the fifth and sixth day of the experiment, with 60.34% females and 39.66% males. The mortality percentage in the different instars for rooms 1, 2 and 4 was: 14.4; 28 and 53.6%, respectively.
Temperature is one of the main environmental variables responsible for changes in the biology of the mosquito A. aegypti, directly influencing the reproductive cycle of this species [27, 28, 29, 30, 31]. The exposure of A. aegypti eggs to temperatures out of their normal range can cause physiological stress, interrupting the development [32]. Therefore, the temperature must be in a suitable range, in order to the larvae hatch and the generations develop and multiply.
In this study, the high hatching rate of larvae (48%) maintained at a temperature of 25°C, in Londrina, Paraná, corroborates what was observed in the laboratory by Farnesi et al. [33] and Mohammed and Chadee [34], who verified larvae hatching rates above 90% at this temperature. In addition, it is in accordance with the optimal temperature range for the development of the A. aegypti mosquito, mentioned in the literature.
Beserra et al. [35] observed that this range is between 21 and 29°C, when studying the thermal requirements of the species in four bioclimatic regions of Paraíba, Brazil. Later, Beserra et al. [36] observed that the optimal temperature for the development of A. aegypti is between 22 and 32°C, when studying the mosquito’s biological cycle in air-conditioned chambers with six different temperatures. For Marinho [37], the ideal temperature range for the development of the vector is between 22°C and 36°C, according to results obtained after analyzing the influence of climatic factors on the pattern of oviposition, distribution and population development of A. aegypti in the field and the laboratory. More recently, Galavíz-Parada et al. [29] conclude that hatching and survival of A. aegypti in Mexico can occur in a temperature range between 15°C and 32°C under laboratory conditions.
Thus, the third highest rate of hatching of larvae (35%), as well as the lowest mortality of larvae (2.2%) occurring at ambient temperature (16.7–24.1°C) in Londrina, is also in line with the range optimal temperature for the development of A. aegypti, mentioned in the literature, and corroborates with a study by Yang et al. [38], where they observed that the mortality rate of adult females of A. aegypti was lower between 15°C and 30°C and increases rapidly at temperatures below or above this range, thus corroborating with the mortality rate observed at 0°C (22.2%) in Londrina. On the other hand, Costa et al. [39] reported that the longevity in A. aegypti decreased from 25 to 35°C, corroborating with the second highest mortality rate, observed at 25° C (16.6%) in Londrina.
Assessing the influence of ambient temperature on the longevity and fertility of A. aegypti in the city of Guarapuava, Paraná, Ajuz et al. [40] found that the vector’s survival zone is wide and that only the minimum temperatures below 5°C limit the proliferation and super infestation of the species in the city. Thus, it is evident that the life cycle of the mosquitoes in Londrina is also affected only in extreme temperatures, in view of the greater number of emergencies of adults of A. aegypti observed at 5°C (44.80%), as well as a relatively low mortality rate on this temperature (6.6%). The ability of A. aegypti larvae and adult to tolerate low temperatures also was demonstrated in the study by Jass et al. [41].
The low hatching of larvae (7%) and emergence of adults (5.6%) at 0°C, as well as the absence at 45°C observed in Londrina, is justified by the fact of extreme temperatures (very low or very high) are harmful to the development of A. aegypti, according to Buriol et al. [42], who stated that temperatures below 5°C and above 40°C are lethal to mosquito development. Similar data were observed by De Majo et al. [43], Marinho et al. [44], Mohammed and Chadee [34] in studies about the effect of different temperatures on the development of A. aegypti. More recently, Sukiato et al. [32] also noted in their study that there was no hatching of A. aegypti eggs at 40°C, and the larval and pupal mortality was higher at 37°C, compared to other lower temperatures (34, 31 and 28°C).
In Manaus, Amazonas, the greatest influence of the room 4 (with the highest temperature - 32.18°C) on the life cycle of the mosquitoes, where they developed faster than mosquitoes kept in other lower temperature environmental rooms, is in line with Carrington et al. [45], who found that temperatures around 30°C are ideal for the development of A. aegypti and that the development was faster under a temperature of 35°C and impaired above this range. Similar results were also obtained by Brady et al. [46], when evaluating the survival of the species at different temperatures, located between 0°C and 40°C. Sukiato et al. [32] also observed that the A. aegypti development time was shorter at higher temperatures (37 and 34°C).
In relation to the mortality rate, the lower mortality (14.4%) that occurred at the room 1, where the temperature was lower, as well as the higher mortality (53.6%) of different instars that occurred at the room 4, also corroborate with the results reported by Yang et al. [38] and Sukiato et al. [32] already compared with the results obtained in Londrina. A study by Farjana et al. [47] also demonstrated that the mortality of A. aegypti increased at a higher temperature (35°C).
The lowest and the highest mortality rate at the rooms 1 and 4, respectively, may have happened due to the influence of CO2 concentration—which was lower at the room 1 (similar to the current atmospheric concentration) and much higher at the room 4—because CO2 atmospheric is also related to the biological cycle of living beings, influencing their ecological interactions. High CO2 rates have an impact on ecological communities, causing a reduction in nitrogen concentration. Furthermore, it can reduce the quality and quantity of food in breeding sites, compromising larval growth and survival [48].
However, in a study by Azevedo et al. [49], higher concentrations of CO2 had no significant influence on the results obtained in relation to the biological cycle of A. aegypti.
In the South region of the country, the hatching of eggs and emergence of adults of A. aegypti only did not occur at 45°C, while the lowest rate of hatching and emergence occurred at 0°C, indicating that the development of the mosquitoes in Londrina is affected only in very extreme temperatures, since the temperature of 5°C still proved beneficial to the development of A. aegypti. In the North region of the country, the development of the immature in Manaus is faster as the temperature in the environmental rooms increases; however, at the same time, death rates also increase.
Therefore, it is concluded that temperatures from 5 to 29.74°C are more appropriate, since values outside these limits can cause deleterious effects in biological aspects related to the reproductive success of the species. Thus, temperature has a great influence on these aspects, with medium temperatures being more beneficial to this species.
The results obtained show that both A. aegypti mosquitoes from the South and North regions of Brazil have adaptive potential in face of the increase in the average temperature stipulated by the IPCC, in view of the unviability of eggs only at extreme temperatures and considering the shorter average duration of the life cycle observed at high temperatures. Thus, the predicted climate changes may favor the development and proliferation of A. aegypti, and consequently the viral circulation, in addition to make possible the occurrence of a geographical expansion of A. aegypti. However, it is important to mention that other environmental variables can also influence the biology of mosquitoes, as well as viruses, requiring, therefore, more studies related to the various environmental variables and viruses in order to be able to affirm whether there will be a greater occurrence of arboviruses due to global warming.
We are grateful to the technicians of the Malaria and Dengue Laboratories (INPA) of the Medical Entomology Laboratory (UEL). This work is supported by the State University of Londrina and ADAPTA/CNPq: 573976/2008-2.
The authors declare no conflict of interest.
Liposomes are artificial lipid-based bilayered vesicles. They were firstly discovered and described in 1965 by Bangham et al. [1] as swollen phospholipid systems, namely Banghasomes. A few years later, the structural description of liposomes was unveiled as small devices made of one or more closed phospholipid bilayers. Due to the diversity of particle sizes, from 20 nanometers to several micrometers, liposomal vesicles are considered as either nanoparticles or microparticles endowed with the ability to encapsulate materials of various nature and polarity [2, 3].
\nUp until now, liposomes have shown huge promise as potential vehicles for biologically active compounds in cosmetic and pharmaceutical industries. These applications have been extended to food and farm industries, where unstable substances such as antioxidants, flavors and antimicrobials have been explored for liposomal encapsulation. Across all these areas of application, liposomes have been deemed to be the most successful delivery systems due to their multiple advantages. These include high biocompatibility and biodegradability, low toxicity and poor immunogenicity, improved drug solubility and controlled distribution, as well as the ability of performing surface modifications for targeted, extended and sustained release. Currently, there are several liposomal formulations that are clinically established for the treatment of various diseases, such as cancer, fungal and viral infections; and many more have reached advanced phases of clinical trials [4, 5].
\nAlthough liposomes have shown some success in drug product development, the limitations identified in liposomal technology have remained almost stagnant over decades. The most common disadvantages of liposomes arise partly from poor stability under shelf and in vivo conditions. This is mostly due to potential lipids oxidation and hydrolysis, leakage and loss of hydrophilic cargoes, as well as particles fission and fusion. To date, some of these problems can be circumvented by playing around formulation adjuvants, such as anti-oxidants, or post-preparation processing, such as freeze-drying [4, 6, 7].
\nWhile describing broadly the current perception of liposomes, regarding production, evaluation and applications; this chapter intends to highlight the longstanding bottlenecks that remained overlooked and challenging to product development and implementation. This would increase the understanding of the gaps in the field and provide future directions to new openings for improvements in liposome technology.
\nThe liposomal vesicles derive from hydration of phospholipids, which are amphiphilic molecules endowed with a hydrophilic head group and two hydrophobic acyl chains (Figure 1). In aqueous media, phospholipid molecules self-assemble into a bilayered structure. Within the bilayer, phospholipid polar groups line up to form a water-attracting surface while their lipophilic chains face each other to yield a water-free zone. On mechanical shaking or heating, phospholipid bilayers continuously enclose the dispersing aqueous medium and form a vesicular system. In this system, hydrophilic groups of phospholipids are oriented towards the inner and outer aqueous phase, while their hydrophobic tails are centered within the bilayer [2, 4]. This architecture underlines the ability of liposomes to readily encapsulate hydrophilic and hydrophobic materials inside the inner aqueous core and the lipid bilayers, respectively (as illustrated in Figure 2).
\nChemical structure of a representative phospholipid molecule (distearoyl phosphatidylcholine).
Flowchart illustrating liposome formation and encapsulation of drug molecules.
Depending on the particle size and number of bilayers forming the vesicles (lamellarity), liposomes can be categorized in the following classes [4, 8]:
Small unilamellar vesicles (SUV), size range 20–100 nm;
Large unilamellar vesicles (LUV), size >100 nm;
Giant unilamellar vesicles (GULV), size >1000 nm;
Oligolamellar vesicles (OLV), size range 100–1000 nm;
Multilamellar large vesicles (MLV), with size >500 nm;
Multivesicular vesicles, size from 1000 nm to several thousand nanometers.
Based on their composition, liposomes can be classified as conventional, long circulating, cationic, stimuli-responsive and immunoliposomes. The differences between these categories will be highlighted later when discussing composition and evolution of liposomes.
\nFurthermore, there are many other vesicular systems considered as part of the liposome-type vesicles. These include emulsomes, enzymosomes, sphyngosomes, transfersomes, ethosomes, pharmacosomes and virosomes, which are lipid-based liposomes analogous. The non-lipid-based liposomes analogous are aquasomes, bilosomes and niosomes [8, 9]. All the liposome-type systems are briefly presented in Table 1.
\nVesicle designation | \nMain components | \nIllustrative application | \n
---|---|---|
Emulsomes | \nA mixture of fats and triglycerides stabilized by high proportion of lecithin | \nEmulsomes loaded with Amphotericin B for the treatment of visceral leishmaniasis [9] | \n
Enzymosomes | \nComplexes of lipids and enzymatic proteins | \nEncapsulation and delivery of superoxide dismutase for oxidative stress management [9] | \n
Sphyngosomes | \nSphingolipids containing amide and ether bonds | \nSphyngosomes loaded with vincristine (Marqibo®) for lymphoblastic leukemia therapy [4] | \n
Transfersomes | \nA mixture of single chain surfactant, phospholipids and ethanol (10%) | \nTransfersomes loaded with diclofenac for improved topical delivery/retention [10] | \n
Ethosomes | \nPhospholipids and ethanol (20–40%) | \nMitoxantrone-loaded ethosomes for the treatment of melanoma [11] | \n
Pharmacosomes | \nConjugate of drug and phospholipid | \nPharmacosomes loaded with diclofenac for enhanced the bioavailability and reduced toxicity [9] | \n
Virosomes | \nViral glycoproteins | \nVirosome containing HIV-1 gp41-subunit antigens for protection against vaginal simian-HIV [12] | \n
Aquasomes | \nTin oxide, diamonds or brushite core covered with oligomeric film | \nPEG-lipid coated aquasomes containing interferon-α-2b for prolonged and enhanced cytotoxicity [13] | \n
Bilosomes | \nBile salts and acids (deoxycholic acid) | \nBilosomes loaded with diphtheria toxoid for systemic and mucosal immunization [14] | \n
Niosomes | \nNon-ionic surfactants (span and tween) | \nNiosomes based formulation for enhanced oral bioavailability of candesartan cilexetil [15] | \n
Presentation of liposome-type systems.
Liposomes are made of physiologically acceptable natural or synthetic phospholipids found in the lipid bilayer membranes of human cells. The most frequently used phospholipids for liposomes preparation are phosphatidylcholines (PC), phosphatidylethanolamines (PE), phosphatidylserines (PS) and phosphatidylglycerols (PG) [16]. The molecular structures of these biocompatible lipids are shown in Figure 3 and Table 2.
\nStructural representation of ester glycerol-phospholipid molecules, with R1 and R2 representing the hydrocarbon chains of different fatty acids.
Description of –X moieties of different glycerol-phospholipids [16].
In liposomal technology, a considerable attention is given to the phase transition temperature (Tt) of these phospholipids. The Tt corresponds to the temperature above which phospholipids exist in liquid crystalline phase. In this fluid state, hydrophobic tails of phospholipids are randomly oriented but ready to form closely continuous bilayered vesicles (liposomes). Below the Tt, phospholipids exist in gel state, where the hydrophobic tails are completely expanded and well packed, thus not able to form liposomes [4, 17].
\nAs most of the phospholipids used for liposomes formulation have Tt close to the physiological temperature (37°C), the addition of cholesterol has been adopted as a strategy to stabilize the liposomal vesicles in physiological media. This is especially for phospholipids that can undergo phase transition and leakage at room temperature, which can lead to premature release of the liposome cargo. In fact, due to its high hydrophobicity, cholesterol was found efficient in strengthening the packing of phospholipid bilayers, reducing therefore membrane permeability. Numerous studies have reported the ability of cholesterol to impact liposomes properties and functionality, including encapsulation efficiency and release characteristics [18, 19, 20]. The work by Kirby et al. [21] demonstrated that increasing cholesterol content can prevent leakage and improve in vivo stability of liposome. Later, Lopez-Pinto et al. [22] observed a direct correlation between cholesterol content and liposome sizes. These observations have established cholesterol content to be a key parameter in liposome formulation.
\nLike cholesterol, there are many other ingredients that can affect liposomes behavior and afford the desired encapsulation or delivery profiles. Additive agents such as oleic acid and N-[1(2,3-dioleoyloxy) propyl]-N,N,N-trimethylammonium chloride (DOTAP) are useful for the preparation of negatively and positively charged liposomes, respectively. These charged liposomes offer the advantage of great liposomal stability during the storage, as charged particles repel each and reduce aggregation tendencies. While the cell internalization of positively charged liposomes (cationic liposomes) is promoted by their electrostatic interaction with cell membranes (which are negatively charged), liposomes bearing negative charges are subjective to poor cell internalization due to the corresponding repulsive forces. Cationic liposomes are used in gene therapy due to their ability to successfully encapsulate nucleic acids by electrostatic forces [4, 23].
\nIn addition, some special lipids such as cholesteryl hemisuccinate (CHEMS) and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), have been widely used to prepare liposomes with pH-dependent release features. CHEMS can exhibit pH-sensitivity either alone or in the presence of other lipids. In ionized forms at basic or neutral pH, CHEMS stabilizes the lamellar form of DOPE in lipid-based vesicles. However, the protonated or molecular CHEMS formed at acidic pH promotes hexagonal phase of this lipid, which leads to the disruption of the vesicular systems and release of the encapsulated materials [24, 25]. Tocopherol hemisuccinate (THS) has also shown similar pH-responsiveness as CHEMS, due to their molecular similarity [26].
\nThe composition of liposome appears to be a broad topic, but also very crucial for the desired product development. However, the nature and costs of the liposome components used over decades viz., particularly the synthetic or highly purified natural phospholipids, have been reported to be part of the factors affecting negatively the universal implementation and affordability of liposome technology [16, 27]. The review by Machado et al. [28] discussed the feasibility of using crude soybean and rice lecithin for liposomal encapsulation of food ingredients. The authors demonstrated these naturally occurring phospholipid mixtures could be useful for liposomes preparation regardless of the intended area of application. Our group has recently investigated liposomal encapsulation of isoniazid using crude soybean lecithin. The formulated liposomes exhibited much better encapsulation efficiency than purified soybean lecithin [29]. This study proposed crude soybean lecithin for liposomal encapsulation of drug molecules. However, the complexity of this lipid mixture might be a bottleneck for some biomedical applications, where molecular architecture of the lipid bilayer is explored to get insights into potential cell membrane permeability. The versatility of crude soybean lecithin liposomes is therefore in question, considering the wide range of areas that the liposomal systems usually cover.
\nBased on the composition, liposomal systems can be considered to have evolved from conventional, long circulating, targeted and immune-liposomes to stimuli-responsive liposomes. The liposomes composed purely of phospholipids with or without cholesterol (conventional liposomes) have shown some limitations due to their uptake by the cells of the mononuclear phagocytic system (MNPS), such as macrophages that ensure liposomes clearance through phagocytosis. This biological fate makes conventional liposomes appropriate vehicles for targeted drug delivery to infected MNPS cells, like the case of alveolar macrophages where Mycobacterium tuberculosis resides often. However, the uptake by the MNPS cells decreases liposomes half-life and exposes to high risk of therapeutic failure when the site of interest is beyond the MNPS [2].
\nExtensive studies conducted in liposome technology led to identification of some astute strategies for addressing the issue of MNPS’ attack viz., liposome downsizing and surface modification. In this regard, it was observed that the physiological clearance of larger liposomes (MLV) was much quicker than that of smaller liposomes (SUL), which describe long-circulating profiles with increased half-lives [30]. The stealth strategy arising from surface modification involved grafting or coating hydrophilic polymers such as polyethylene glycol (PEG) and chitosan, which prevents detection of liposomes by the MNPS cells. While stealth behavior allows liposomes to achieve much longer circulation time (hence the name “long circulating liposomes”), this strategy comes along with poor targeting efficiency due to wider distribution of liposomes in the body. From this limitation, further developments have led to introduction of targeted liposomes. These liposomes are characterized by surface decoration with glycoproteins, polysaccharides or specific receptors ligands to achieve narrowed distribution and accumulation at the site of interest [9, 31]. The observation that ligand-decorated liposomes could provide selective drug accumulation inspired further design of antibody-functionalized liposomes (immunoliposomes) as well as stimuli-responsive liposomal systems for controlled drug delivery [32]. Figure 4 shows the trend in the development of “intelligent” liposomes for site-specific delivery; from conventional liposomes, stealth liposome, targeted liposomes, immunoliposomes to stimuli-responsive liposomes.
\nSchematic representation of the trend in liposome improvements for site specific delivery. (A) Conventional liposomes, (B) stealth liposome, (C) targeted liposomes, (D) immunoliposomes and (E) stimuli-responsive liposomes [32].
Stimuli-responsive liposomes are smart liposomal systems that exhibit rapid release of the cargo upon changes in some physicochemical or biochemical stimuli, such as pH, temperature, redox potentials, enzymes concentrations, ultrasound, electric or magnetic fields [33, 34]. Among these stimuli, pH change is the most promising stimulus due to the existence of multiple pH gradients in the body [35]. In common practice, formulation of pH-sensitive liposomes involves incorporation of CHEMS and DOPE. Although CHEMS-DOPE-based liposomes have shown great promise for controlled delivery [24, 25, 36], the costly status of these lipids remains a deep concern for wider development and application of liposome products. To circumvent the use of such special lipids for pH-dependent delivery, our group has considered hydrazone derivatization of isoniazid (INH), as a small hydrophilic drug model, followed by encapsulation using crude soybean lecithin for cost-effective development [37]. Being poorly water soluble, the prepared conjugates were successfully embedded within the lipid membranes and INH release experiments were conducted in different pH media. The INH-conjugate loaded crude soybean lecithin liposomes have demonstrated attractive nanoparticulate and stimuli-responsive characteristics for potential low-cost site-specific liposomal delivery [38]. However, the amounts of INH-derivatives loaded were found to be almost 10-folds lesser than the loading achieved when native INH was encapsulated in crude soybean lecithin liposomes, which was in turn associated with some burst release [29]. This is probably due to the fact that INH derivatives are encapsulated in the lipid bilayer, which offers limited space for loading [39], while the native INH was trapped within the aqueous core of liposomes. These limitations underline the need for further developments in liposome technology to achieve controlled release from cost-effective liposomes, combining both use of cheap lipids and high drug loadings.
\nLiposomes can be prepared using a wide range of methods that involve combination of lipids with aqueous media, and somehow affect liposomes characteristics, such as size, lamellarity and encapsulation efficiency (EE). The recently reported methods can be categorized as conventional, which mostly involve approaches that are easy to use at laboratory scale, and novel methods that appear to be more useful for up-scale production but require some special equipment [4].
\nThe most commonly used methods for formulation of liposomes share the following fundamental stages: (i) lipids dissolution in organic solvents, (ii) drying of the resultant solution, (iii) hydration of dried lipid (using various aqueous media), (iv) isolation of the liposomal vesicles, and (v) quality control assays [6]. While sharing these basic stages, the conventional preparation methods gather different advantages and disadvantages that are comparatively presented in Table 3. The specific technological details of these methods are separately discussed in the following paragraphs.
\nMethod designation | \nAdvantages | \nDisadvantages | \n
---|---|---|
Film hydration (Bangham method) | \nStraightforward process | \nUse of organic solvent and mechanical agitation, production of large particles with no control on size, poor encapsulation efficiencies of hydrophilic materials, time consuming, sterilization issue | \n
Reverse phase evaporation | \nSimple design, suitable encapsulation efficiency | \nNot applicable to fragile cargoes, use of large quantity of organic solvent, time consuming, sterilization issue | \n
Solvent injection | \nStraightforward approach | \nTrace of organic solvent as residue, possible nozzle blockage in ether system, time consuming, sterilization issue | \n
Detergent removal | \nSimple design, homogenous product, control of particle size | \nPresence of organic solvent, detergent residue, time consuming, low entrapment efficiency, poor yield, sterilization required | \n
Heating method | \nSimple and fast process, organic solvent free, no need for sterilization, possible up-scale production | \nThe need for high temperature | \n
Also known Bangham method, film hydration represents the simplest and oldest method used in liposome technology. In this method, lipids are firstly dissolved in a suitable organic solvent, and dried down to yield a thin film at the bottom of the flask. The obtained lipid film is hydrated using an appropriate aqueous medium to produce liposomal dispersion. The structural organization of the formed vesicles can be affected by the hydration conditions. A gentle hydration of the lipid film forms giant unilamellar vesicles (GULV), whereas a hash hydration gives rise to multilamellar vesicles (MLV) with poor size homogeneity, which requires an additional downsizing step. The most commonly used sizing methods include probe and bath sonication that afford production of small unilamellar vesicles (SUV). Despite its higher effectiveness, probe sonication is often blamed for potential contamination (with titanium from the titanium-based nozzle used for mechanical agitation), and production of local heat that can affect lipids and drugs stability. Although the two sonication methods produce liposomes with identical characteristics, the use of bath sonication remains a better option due to easy control of operational parameters. Another technique used for liposome sizing includes consecutive extrusion of the liposomal formulation through polycarbonate filters of defined pore sizes. In this method, the number of extrusion cycles is the key parameter to control for effective homogenization [4, 6].
\nReverse phase evaporation is an alternative method to the film hydration that involves formation of water-in-oil emulsion between the aqueous phase (containing hydrophilic materials) and the organic phase (containing lipids and any hydrophobic materials). A brief sonication of this mixture is required for system homogenization. The removal of the organic phase under reduced pressure yields a milky gel that turns subsequently into liposomal suspension. The liposomes can be isolated from the dispersion using centrifugation, dialysis or sepharose 24 column [28].
\nSolvent injection involves quick injection of the lipid solution (in ethanol or diethyl ether) into an aqueous medium. The experiment is performed either at room or at higher temperature (e.g., 60°C), depending on whether the organic solvent is water-miscible or not. The liposomes prepared by solvent injection process are mostly polydispersed and highly contaminated by organic solvents, especially ethanol due to formation of azeotrope mixture with water. As presented in Table 3, solvent injection suffers from several drawbacks including continuous exposure of the therapeutic agents to high temperature and organic solvent that might affect both the stability and safety of the liposomal products [28, 40].
\nIn the detergent removal method, phospholipids are dissolved in aqueous solution containing detergents at critical micelle concentrations (CMC). Upon detergent removal, the reaction medium frees individual phospholipid molecules that self-assemble into bilayered structures. Detergent removal is mostly achieved by means of a dialysis bag, polystyrene-based absorber beads or Sephadex columns (gel permeation chromatography). Dilution of the resultant mixture with some appropriate aqueous medium leads to restructuration of the formed micelles that evolve to liposomes [4, 6].
\nAmong all the conventional methods, the heating method is known to be the most attractive method for liposomes preparation due to its organic solvent free characteristics. In the heating method, lipids are hydrated for 1 hour, and heated for another hour above the transition temperature of the phospholipids in the presence of a hydrating agent (glycerin or propylene glycol 3%). When cholesterol is part of the formulation, the reaction medium is heated up to 100°C because of its high melting point. Being prepared under heating conditions, the resultant liposomes can be readily used without any further sterilization treatments, which minimizes both formulation complexity and timing. In addition, there is no need for further removal of the hydrating agents employed, since these represent physiologically acceptable ingredients that are well-established for pharmaceutical applications. Moreover, the observation that these hydrating agents can prevent particle coagulation and sedimentation makes them much more attractive as stabilizer and isotonizing additives. The hydroxyl groups of these hydrating agents provide a cryoprotective effect that makes the heating method an efficient method for the formulation of inhalable liposomes [41, 42].
\nMicrofluidic methods include all the novel techniques that make use of microscopic channels (in the size range of 5–500 μm). In this method, lipids are dissolved in an appropriate organic solvent (ethanol or isopropanol) and the resultant solution is propelled perpendicularly or in the opposite direction to the aqueous medium within the micro-channels. The continuous axial mixing of the organic and aqueous solutions leads to liposomes formation because of local diffusion of phospholipids in aqueous phase, which encourages the self-assembly process. Among many others, the micro hydrodynamic focusing method represents the most commonly used microfluidic method for liposomes formulation. This method produces small and large unilamellar vesicles, 40–140 nm, with good size homogeneity (mono dispersed feature). The other microfluidic techniques include the microfluidic droplets and the pulsed jet flow microfluidic methods. The microfluidic droplets method involves dissolution of phospholipids in hexane for preparation of giant liposomes (4–20 μm). In the pulsed jet flow microfluidic method, the conventional film hydration method has been modified by drying the lipid solution in microtubes. The resultant lipid film is hydrated within the microtubes through a perfusion process that produces much larger vesicles, 200–534 μm, with remarkable encapsulation efficiency [4, 43]. As common advantages, the microfluidic methods offer the possibility for production of vesicles with desired size, due to the versatility and flexibility of the methods. The disadvantages of these methods include the imperative use of organic solvent and drastic agitation, as well as difficulty for large scale production [40].
\nWhile being considered as equivalent to the conventional reverse phase evaporation method, supercritical fluidic technique represents the most important novel liposome preparation methods that makes use of a supercritical fluid, such as carbon dioxide (CO2) maintained under supercritical conditions (supercritical temperature and pressure). In this state, CO2 is an excellent solvent for the lipids. The high-performance liquid chromatography (HPLC) pump provides a continuous flow of the aqueous phase into a view cell that contains the supercritical lipid solution, allowing phase transition of the dissolved phospholipids. Upon sudden decrease in pressure, CO2 gets completely removed and phospholipids self-assemble into a bilayered vesicular system. The supercritical fluidic method affords large unilamellar vesicles (100–1200 nm) with 5-fold higher encapsulation efficiency than the equivalent conventional method. Apart from being organic solvent-free methods, the supercritical fluidic method offers many other advantages such as the use of CO2, as a cheap and environmentally harmless solvent, possibility for controlling particle size, in situ sterilization and large-scale production in industrial settings. However, the disadvantages of the supercritical fluidic technique, including particularly its high cost, low yield and use of high pressures (200–350 bar) which require special infrastructures, restrict their universal applications for wider developments of liposomal technology [4, 40, 43].
\nThe freeze-thawing treatment involves freezing the liposomes dispersion in liquid nitrogen, and subsequently thawing it at the temperature above the phase transition temperature of the lipids used for formulation. Upon freeze-thawing, the liposomal vesicles are subjected to fusion since the lipid bilayers become fluid and highly permeable, allowing extensive diffusion of hydrophilic molecules, which leads to important cryoconcentration. These structural modifications encourage encapsulation of hydrophilic materials that are poorly loaded in liposomes when conventional methods are used. This underlines the reason why freeze-thawing represents an important treatment in liposome technology. Amongst the key parameters to be considered for freeze-thawing optimization are the number and duration of freeze-thawing cycles. These can impact significantly not only the encapsulation efficiency but also structural characteristics, i.e., liposomes lamellarity and polydispersity [44, 45].
\nCommonly known as lyophilization, freeze-drying is a post-preparation treatment for liposomes that is applied in both laboratory and industrial settings to preserve the characteristics of liposomal products. Freeze-drying involves freezing of the aqueous samples and subsequent removal of ice by sublimation. Freeze-drying represents a very useful treatment for shelf stability of liposomal suspensions, since water molecules can trigger some chemical reactions and lead to modification of the cargo or excipients in the formulation. Freeze-drying appears to be of great interest when the prepared formulation contains thermos-sensitive materials such as proteins, nucleic acids, etc., which might undergo fast degradation when subjected to heat-drying. The use of freeze-drying has gained considerable attention in liposome technology due to improved storage stability of liposomal products. Because of potential leakage of liposomes during freeze-drying, addition of hydrophilic compounds, commonly called cryoprotective agents (such as carbohydrates), has been established to ensure good stability and quality of the final product. The cryoprotectants commonly used include mannitol, lactose, sucrose and trehalose. Among these, trehalose is the most reputed cryoprotecting agent since it preserves liposomes stability during and after freeze-drying treatment [6].
\nAfter production, liposome formulations are subjected to extensive characterization, evaluating the physicochemical properties of liposomes that affect their shelf stability and biological performance. The most routinely investigated parameters in liposome characterization include vesicle size and size distribution (or polydispersity), surface charge (or Zeta potential), shape and morphology, lamellarity, encapsulation efficiency, phase behavior (or polymorphism) and in vitro release profile [4, 46]. Table 4 indicates the techniques used for evaluation of liposome characteristics. The most frequently used methods are briefly discussed in the following paragraphs.
\nParameters | \nAnalytical techniques | \n
---|---|
Particle size | \nDynamic light scattering (DLS), size exclusion chromatography (SEC), field-flow fractionation (FFF) and microscope technology: transmission electron microscopy (TEM), cryogenic-TEM (Cryo-TEM), and atomic force microscopy (AFM) | \n
Zeta potential /Surface charge | \nElectrophoretic mobility, DLS | \n
Particle shape /morphology | \nMicroscopic techniques such as TEM, Cryo-TEM and AFM | \n
Lamellarity | \nCryo-TEM and 31P-NMR | \n
Phase behavior | \nX-ray diffraction (XRD), differential scanning calorimetry (DSC) and thermogravimetric analysis (TGA) | \n
Encapsulation efficiency | \nCentrifugation, dialysis or column separation for liposomes isolation, followed by drug content determination | \n
Drug release | \nDialysis or centrifugation, followed by drug quantification using analytical method, such as UV–Vis spectrophotometry, fluorescence spectrometry, enzyme- or protein-based assays, gel electrophoresis, HPLC, UPLC, LC-MS | \n
Analytical methods commonly used for liposomes characterization.
Also known as photon correlation spectroscopy or quasi-elastic light scattering, DLS represents the most commonly used method for determination of liposome size, size distribution (polydispersity) and Zeta potential (surface charge). DLS is done by an instrumental setting called Zetasizer Nano. The standard operational principle of DLS is based on continuous motion of dispersed particles due to their bombardment by solvent molecules (Brownian motion). This phenomenon causes remarkable scattering of the applied light. Since the extent of fluctuation in light intensity is associated with the diffusion rate of the suspended particles, which is related to particle diameter (smaller particles diffusing faster than the larger ones), the particle size is automatically deducted from the estimated amount of the scattered light. When addressing Zeta potential measurements, DLS allows surface charge determination by accessing changes in the scattered light intensity caused by particle motion due to the electric field applied. In other words, for surface charge (Zeta potential) evaluation, changes in the intensity of the scattered light are governed by the applied electric field (which causes extensive motion of charged particles), in contrast to size measurements where Brownian motion is the key factor [4, 47]. Apart from being a simple, fast and reliable method for routine analyses, DLS offers many other advantages including the fact that the measurement is taken from a native environment, and a wide size range can be evaluated (from a few nanometers to several micrometers). However, DLS shows some limitations such as the difficulty of differentiating individual particles from aggregates and high sensitivity to contaminants [48]. In addition, DLS is technically unable to provide true particle size, but rather hydrodynamic diameter due to particle solvation. Water layers on particle surface lead to false readings of particle diameters in aqueous media [49].
\nThe microscopic observation provides direct visualization of the liposomal vesicles as individual particles, which allows effective analysis of shape and morphology as well as a precise and reliable size reading. In this context, TEM techniques are commonly used in liposome technology for structural characterizations. In TEM experiments, the liquid sample is spotted onto a copper grid, and the solvent dried prior to the microscopic analysis. Under TEM instrument, liposomal vesicles mostly appear as black spherical particles on a white background. For a variant TEM technique like negative staining TEM, liposomes appear as bright spherical spots on a black background since the spotted sample is treated with uranyl acetate or phosphotungstic acid (as negative staining agent). Due to its effectiveness, TEM appears to be a powerful complementary technique to DLS for confirmation of the liposomal structure. Unlike DLS, TEM offers the advantage of differentiating individual vesicles from aggregates, allowing critical assessment of the liposome population. Nevertheless, TEM presents several limitations due to sample preparation. Apart from being time-consuming, sample pretreatments in TEM analyses may cause remarkable changes in liposomal shape or morphology: potential vesicle shrinkage, swelling or artifact formation [4, 50].
\nTo overcome these limitations, Cryo-TEM was developed as a strategy to minimize liposome disruption by making use of a flash freezing treatment for direct particle visualization in solid-state (without solvent removal). Nowadays, Cryo-TEM is the most reliable technique for visual determination of liposome structure including lamellarity. However, Cryo-TEM appears to be sometimes limited since it works perfectly only with very small particles. This has led to the development of atomic force microscopy (AFM) for direct particle analysis in native environments. Although AFM offers the advantage of higher particle resolution at three-dimensional level, the use of this technique is mostly limited by the high cost of the instrument, which compromises its universal availability and accessibility [3, 4].
\nDSC and XRD are complementary techniques that evaluate the thermal behavior and crystallinity, respectively, and provide valuable information for characterization of loaded liposomes [46, 51, 52]. DSC evaluates the differences in heat flow (electric power) between a sample and a reference. In DSC experiments, the sample and the reference are subjected to a programmed heating, cooling or isothermal treatment in a controlled atmosphere (mostly saturated with nitrogen gas). The heating treatment is achieved either by the same heater (heat flux DSC) or by separate heaters (power compensated DSC). The experiment is conducted in specialized metal pans made of aluminum, tin, zinc or indium. Throughout the experiment, frequent electric power adjustments occur upon material phase transition (melting or crystallization), ensuring thermal equilibrium between the sample and the reference. This phenomenon is described and expressed by the plot of heat capacity against temperature or time (heat flow curve). The heat flow curve provides the respective transition temperature and enthalpy, which allows to identify the nature of thermal events: endo- or exothermic [53, 54, 55]. DSC represents the most useful thermal analysis technique in the study of lipid-based materials [46, 52, 54, 55, 56].
\nUnlike DSC where sample recovery is not possible, XRD is a non-destructive analytical tool that allows structural investigations of crystalline materials. XRD makes use of X-rays that deeply penetrate solid materials and provide useful information at atomic structure level. Though relatively expensive, an XRD instrument is an environmentally and user-friendly device that is easy to use. A wide range of materials such as powders, crystals and liquids can be quickly assessed by XRD. Its other advantages include high resolution, reliability, relatively cheap maintenance, and easy data collection, processing and interpretation. The phase transitions and polymorphism determined by XRD represent valuable information in pharmaceutical development and production of both excipients and biologically active materials [57, 58, 59].
\nThe lamellarity of liposomes is part of their structural characteristics that can have an impact on the intended applications. The number of lipid bilayers can be evaluated using chemically labeled or radiolabelled agents that can be distributed in the bilayer membranes. However, this technique is limited since these reagents might be distributed only on the outer lipid membrane and lead to false readings. To date, Cryo-TEM is the most commonly used technique for the determination of lamellarity by visualization [4]. The nuclear magnetic resonance spectroscopy of the 31-phosphorus (31P-NMR) is also being currently used to estimate the lamellarity of liposomes. This technique deals with the estimation of the ratio of phospholipid amount in the outer layers to that of the inner layers [60]. The 31P-NMR spectrum with a broad peak indicates the presence of MLV while a narrow peak corresponds to SUV. The addition of paramagnetic ions such as Mn2+, Co2+, and Pr3+ shifts the respective peaks to either downfield or upfield because of ionic interactions with the phosphate backbone. By comparing the spectroscopic profile with and without the paramagnetic ion, the lamellarity of liposomes can be estimated. Some other techniques such as small-angle X-ray scattering (SAXS) and trapped volume determination can be used to estimate liposome lamellarity [4, 61].
\nThe profile of release for the liposomal cargoes is commonly estimated in vitro using dialysis. This method implies trapping the liposomal dispersion into a dialysis bag of specific molecular weight cut off. The resultant tubing membrane is placed in a simulated physiological fluid (release medium) that is often a buffer maintained under well-defined conditions: specific temperature and speed of stirring/shaking. At predetermined time intervals, an aliquot is withdrawn from the release medium and an equal volume of the fresh buffer is replaced to maintain sink conditions. In the withdrawn sample aliquots, the released cargo is quantified using some routine analytical techniques such as UV-Vis spectrophotometry, HPLC, UPLC, etc., adapted to the molecular species under evaluation. The release profile is obtained by plotting the cumulative release percentage against the chosen time intervals [4]. Data from the in vitro release study are valuably considered as part of the rational development of formulations for controlled release, since they allow effective prediction of in vivo performance of the delivery systems [62].
\nLiposomes have evolved so far from mere experimental tools of research to industrially established products for clinical and veterinary use. They have shown the ability to improve the physicochemical features of the cargoes and ferry them to the sites of interest. The concepts of liposomal encapsulation have been applied in several fields of life science. Liposomes are frequently used for the delivery of drug, gene, vaccine and diagnostic products; but other applications encompass encapsulation of food and cosmetic ingredients as well as routine analysis of chemical substances [2, 3]. The following paragraphs briefly present the current applications of liposomes.
\nThe use of liposomes in drug delivery aims at modifying the pharmacokinetics of drugs to improve the therapeutic efficacy while minimizing potential toxicity [6]. Liposomes can alter the spatial and temporal distribution of the entrapped drug molecules in vivo, leading to controlled delivery at the site of interest and reduced off-target adverse effects [63]. The liposomal systems have been extensively investigated for the delivery of existing and emerging drugs at various research levels, from basic stages related to research and development to preclinical and clinical applications. Nowadays, liposomes represent the most clinically established drug vehicles for human diseases [3, 5]. The efforts invested in liposomal technology have so far led to the development of several effective liposomal formulations that are currently used in clinics (Table 5).
\nBranded product | \nDrug name | \nTherapeutic indications | \n
---|---|---|
Abelcet | \nAmphotericin B | \nFungal infections | \n
AmBisome | \nAmphotericin B | \nFungal infections | \n
Amphocil | \nAmphotericin B | \nFungal infections | \n
DaunoXome | \nDaunorubicin | \nHematological malignancy | \n
DepoCyt | \nCytarabine | \nLymphomatous meningitis | \n
DepoDur | \nMorphine sulfate | \nPain relief | \n
Doxil | \nDoxorubicin | \nKaposi’s sarcoma and solid tumors | \n
Epaxal | \nInactivated hepatitis A virus | \nHepatitis A | \n
Evacet | \nDoxorubicin | \nOvarian cancer | \n
Inflexal V | \nInactivated hemagglutinin of influenza virus strains A and B | \nInfluenza | \n
LipoDox | \nDoxorubicin | \nKaposi’s sarcoma and solid tumors | \n
Marqibo | \nVincristine sulfate | \nAcute lymphoblastic leukemia | \n
Visudyne | \nVerteporfin | \nPhotodynamic therapy | \n
Liposomal formulations have been used to address a wide range of pathological conditions through different administration routes, including dermal, transdermal, oral, pulmonary and parenteral routes. The clinical areas commonly explored in liposome research encompass skin disorders, cancers and infectious diseases [4]. Amongst, cancer therapy appears to be in the forefront of liposome delivery, due to poor bioavailability and side effects of most of the anti-cancer drugs. However, several infectious diseases, most specially where the pathogen is hosted by the MNPS (i.e., tuberculosis, leishmaniasis, fungal infections), have been reported to be good candidates for liposome application, taking advantage of the spontaneous liposomes uptake by the cells of MNPS [2]. Apart from the nature and localization of the disease, the design and development of liposomes depend also on the intended administration route, since different anatomical and physiological characteristics can be encountered from one route to another [46].
\nFurthermore, the application of liposomes in drug delivery is highly dependent on their colloidal and physiochemical features, i.e., vesicle size, surface charge and system stability [64]. For instance, small liposomes (SUV) are good candidates for Parkinson’s and Alzheimer’s diseases, due to the need for crossing the brain blood barrier to achieve brain targeted delivery. Meanwhile, large liposomes are preferred for macrophage targeted delivery of antimicrobials, when pathogens are located inside the MNPS cells (e.g., tuberculosis, leishmaniasis), taking advantage of the passive liposome cell uptake [2, 4, 46]. This underlines the need for thorough exploration of process and formulation parameters at early stages of products development to produce liposomes with desired characteristics, making the technology for liposomes manufacturing key to future therapeutic research and development.
\nLiposomes have been reported to achieve effective intracellular delivery of genes. These liposomes, also called lipoplexes, are generally made of cationic lipids, which allow for encapsulation of genetic materials via electrostatic interactions with the negatively charged phosphate backbones of nucleic acids. The positive charge on the surface of liposomes also influences their interactions with negatively charged cell membranes and promotes cell internalization [23, 65]. Cationic liposomes offer several advantages over viral gene vectors, including easy and safe production, cost effectiveness, possibility of monitoring toxicity, biodegradability, biocompatibility and lack of dangerous immunogenicity. However, they are suffering from poor transfection efficiency due to their limited endosomal escaping ability, which exposes the genetic materials to enzymatic and acid degradation in lysosomes. This has been improved by incorporating fusogenic lipids, such as DOPE, in the liposomal formulation. These helper lipids facilitate endosomal escaping by membrane fusion with endosomes, leading to early cytoplasmic release of the gene. Though cationic liposomes have shown some dose-dependent toxicity, successful results have been obtained in cancer therapy when delivering genes encoding for tumor suppression proteins [66, 67]. He et al. have recently developed folate receptor alpha-targeted lipoplexes with therapeutic gene expression regulated by an hTERT promoter. These liposomes have shown some promise for the treatment of ovarian cancer [68].
\nThe use of specific lipids or molecules such as phosphatidylserine, DOTAP, fatty acids and monophosphoryl lipids can produce liposomes with attractive immune-stimulating activities. Liposomes carrying antigenic materials, either encapsulated in the aqueous core, grafted or coated on the surface; can stimulate immune responses on macrophage uptake. Following endolysosomal degradation, macrophages present the antigen to T-lymphocytes that initiate the production of cytokines and specific antibodies via activation of B-lymphocytes [4]. The immune response produced by liposomes can be influenced by their composition, lamellarity, size and surface charge [69]. Liposomes containing a glycolipid, trehalose 6,6\'-dibehenate, and a cationic lipid, dimethyldioctadecylammonium, in a 1:5 mass ratio have demonstrated efficient delivery of the TB vaccine Ag85B-ESAT-6. This vaccine has shown prolonged immune response without any toxic effects [70]. Although a liposomal vaccine (namely Stimuvax®) targeting the major histocompatibility class I complex for lung carcinoma was not successfully implemented, some other vaccines such as Epaxal®, Inflexal® V and Mosquirix® have been clinically established for the treatment of hepatitis A, influenza virus infections and malaria, respectively. These vaccines, classified as virosomes, are liposomes generally made of reconstituted viral envelop supplemented with phosphatidylcholine. They offer the advantage of undergoing membrane fusion either with the cells or the endosomes, and thus leading to efficient cytosolic delivery [2, 9].
\nThe use of liposomes for diagnostic purposes is one of current topics of great interest in biomedical applications. Liposomes with magnetic properties, also called magnetoliposomes, are made by entrapping superparamagnetic iron-based nanoparticles or iron oxides or gadolinium (III) chelates for magnetic resonance imaging. Functionalized liposomes labeled with radioisotopes can also be used as molecular probes in nuclear imaging. For instance, liposomes labeled with 64Cu, 18F, 89Zr or 52Mn have been reported in positron emission tomography while 99mTC, 111In or 67Ga labeled liposomes were applied in single photon emission computed tomography. Acoustic liposomes which are liposomes made of perfluoropropane gas can be used as contrast agents in ultrasound imaging technique. The encapsulation of quantum dots and fluorescent dyes into liposomes has also led to the development of attractive liposomal platforms for diagnosis. Additionally, liposomes have shown great potential for simultaneous accommodation of drugs and diagnostic agents such as radionuclides, magnetic or contrast substances. This can be achieved by encapsulation in the inner core, embedding in the lipid bilayer, chemically grafting or coating onto the surface of liposomes. A successful targeted co-delivery of these materials has given to liposomes the status of theranostic systems, as they provide both the therapeutic effect and the diagnostic control [2, 71, 72].
\nIn analytical domains, liposome-based formulations can be usefully involved in immunoassays, biosensors analysis and liquid chromatography. Liposomes can be used to encapsulate, embed or conjugate the analytical entities with high loading capacity and huge surface area that can enhance the intensity of analytical signal. In a direct enzyme-linked immunosorbent assay (ELISA) like method, fluorophore-conjugated liposomes carrying a substantial amount of secondary antibody quickly bind to the antigen that has been fixed on the primary antibody. The addition of a colorimetric substrate leads to color development and allows for analytical estimation of the antigenic analyte. The use of liposomes has been deemed to lower the limits of detection of analytes and increase the sensitivity of immunological analysis technique [73]. In chromatography, the conjugation of liposomes to the stationary phase for gel permeation chromatography is useful for separation of drugs and proteins and for exploring possible molecular interactions on phospholipid membranes. Additionally, the cell-like appearance of liposomes makes them appropriate simulated cell models for studying and predicting the interactions between biologically active compounds and cell membranes [4].
\nThe use of liposomes in the field of cosmetics is based on the similarity between the lipid composition of the liposomal vesicle and that of the biological layers composing the skin. Because phospholipids can be subject to hydration, topical liposomes happen to contribute to the reduction of skin dryness. While playing the role of attractive vehicles for relevant cosmetic agents, liposomes provide a great source of skin ingredients, such as essential unsaturated fatty acids like linolenic acid. This has shown great potential for maintaining the skin and hair in good physiological standing, preventing the rise of some common topical diseases. Furthermore, the use of skin care formulations made of empty or hydrating agents loaded liposomes helps to reduce the transdermal water loss, which is a major cause of skin dryness [74, 75].
\nAs versatile lipid-based systems, liposomes have shown some potential in the encapsulation of food ingredients. In this field, the use of liposomes aims to stabilize some nutraceutical or dietetic ingredients during the storage, to improve their organoleptic characters or to provide a controlled and targeted delivery of these substances in a specific tissue. The shelf life and efficacy of instable bioactive products such as vitamins, enzymes and anti-oxidative agents have been prolonged by their liposomal encapsulation. For example, proteinase and lipase loaded liposomes have been reported to improve the ripening of cheese notably in preventing proteolysis of casein [28, 76]. Yokota et al. [27] have successfully improved the taste and odor of casein hydrolysate in dietetic preparations by liposomal encapsulation. In addition to the encapsulation of dietetic compounds, liposomes have been recently used to encapsulate a cyanobacterium, namely Spirulina platensis, as a source of proteins [77]. Although the use of liposomes in food industry is still at the infant stages of development, advanced studies have been launched to investigate much more attractive applications, such as detection of food contaminants by means of stimuli-responsive liposomes for food safety [76].
\nLiposomes appear to be reputed carriers for various chemical and macromolecular species. From way back their discovery, liposomes have been subject to extensive evolution, in terms of composition, manufacturing and usages, which led to several openings in both basic and applied life sciences. The general details presented herein attempted to reveal some of the existing gaps in liposomes technology and open new windows for further research. An important future breakthrough could be the discovery of cost-effective materials to formulate liposome vesicles with remarkable versatility viz., being suitable as vehicles for various molecules while holding a clear molecular architecture to act as cell models for in vitro bioassays. In addition, novel manufacturing methods for facile encapsulation of both hydrophobic and hydrophilic molecules with no need for organic solvents and special equipment or sophisticated infrastructures are highly desired. Furthermore, the critical issue of system instability for liposomes loaded with hydrophilic materials requires new strategies that will achieve acceptable loading while aiming at targeting cargo delivery at the site of interest. Finally, since liposomes have demonstrated some clinical success as drug vehicles, future efforts should be dedicated to ensuring wider developments and implementation of therapeutic liposomes. This would enhance commercial availability and accessibility of liposome products across the globe, particularly in low- and middle-income countries.
\nWe acknowledge the support received from the National Research Foundation of South Africa and the Sandisa Imbewu Programme of Rhodes University. CI Nkanga and AM Bapolisi are grateful to the NGO Förderverein Uni Kinshasa e. V.-BEBUC/Else-KroenerFresenius Stiftung and Holger-Poehlmann for the advice.
\nThere is no conflict of interest to declare.
"I work with IntechOpen for a number of reasons: their professionalism, their mission in support of Open Access publishing, and the quality of their peer-reviewed publications, but also because they believe in equality. Throughout the world, we are seeing progress in attracting, retaining, and promoting women in STEMM. IntechOpen are certainly supporting this work globally by empowering all scientists and ensuring that women are encouraged and enabled to publish and take leading roles within the scientific community." Dr. Catrin Rutland, University of Nottingham, UK
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