von mardiyanto saarbrücken 2013

Investigation of Nanoparticulate Formulation Intended for Caffeine

Delivery to Hair Follicles

Dissertation

zur Erlangung des Grades

Doktor der Naturwissenschaften

der Naturwissenschaflich-Technischen Fakultät III

Chemie, Pharmazie, Bio-und Werkstoffwissenschaften

der Universität des Saarlandes

von

Mardiyanto

Saarbrücken

2013

Tag des Kolloqiums :

Dekan : Prof. Dr. Volkhard Helms

Berichterstatter : Prof. Dr. Marc Schneider (Universität des Saarlandes)

Prof. Dr. Alexandra Kiemer (Universität des Saarlandes)

Vorsitz : Prof. Dr. Gregor Jung (Universität des Saarlandes)

Akad. Mitarbeiter : Dr. Maike Windbergs (Universität des Saarlandes)

16. 07. 2013

Die vorliegende Dissertation entstand unter der Betreung von

Prof. Dr. Marc Schneider

in der Fachrichtung Biopharmazie und Pharmazeutische Technologie

Arbeitsgruppe Pharmazeutische Nanotechnologie

der Universität des Saarlandes

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Table of Contents

Table of Contents ........................................................................................................

Short Summary ...........................................................................................................

1

4

Kurzzusammenfassung ..............................................................................................

5

1. Introduction ...........................................................................................................

6

1.1 Hair loss problem in human life .......................................................................... 6

1.1.1 Internal and external factors causing hair loss ………………………………… 6

1.1.2 Prevention and treatment of hair loss …………………………………………... 7

1.1.3 Caffeine as stimulating agent to proliferate hair growth ………….………..….

8

1.2 Topical and follicular nanoscale drug delivery syst ems …………….…………. 9

1.2.1 Topical nanoscale drug delivery systems ………………..…………………….. 10

1.2.2 Penetration pathway across the skin …………….…….……………………….. 12

1.2.3 Transfollicular pathway for drug delivery systems …………….….…………...

13

1.3 Particulate formulation nanoscale drug delivery sys tems …………..….……... 16

1.3.1 Current status of polymeric nanoparticles ……………………………………… 16

1.3.2 Preparation of polymeric nanoparticles ………………………………………… 18

1.3.3 Chitosan and chitosan-PLGA nanoparticles …………………………………… 20

1.3.4 Magnetite loaded polymeric nanoparticles …………………..….……………...

21

2. Aim of dissertation and experimental design ……………….…………………….

23

3. Interaction of chitosan and caffeine …………………….…………………………..

26

3.1 Introduction ………………………………………..…………………….………………

26

3.2 Materials and methods …………………………………..…………….……………… 27

3.2.1 Materials ……………………………………..…………………….………………. 27

3.2.2 Equipment …………………………………………..……………….…………….. 27

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3.3.3 FTIR measurement ……………………………………………………………….. 27

3.2.4 DSC measurement ……………………………………..…………….…………... 27

3.2.5 Determination of chitosan and caffeine by UV spectroscopy ……...………… 28

3.2.6 Solubilization study of caffeine in the presence of chitosan ……..….……….

28

3.3 Results and discussion ……………………………..………….……………………..

29

3.4 Conclusions ……………………………………………………………………………..

34

4. Nanoparticulate formulation intended for hair fo llicle targeting ………...…….

36

4.1 Introduction …………………………………..………………………………….………

36

4.2 Materials and methods …………………………………..………………….………… 38

4.2.1 Materials ……………………………………..…………………………….………. 38

4.2.2 Equipment …………………………………………………..……………….…….. 38

4.2.3 Preparation of Nanoparticles ………………………………………..…….…….. 38

4.2.4 Purification and %EE of caffeine ……………………………………..…….…… 39

4.2.5 Size, PDI, zeta potential and morphology of nanoparticles …………….…..... 40

4.2.6 Dissolving nanoparticles and determination of caffeine loading ………...…... 40

4.2.7 In vitro release study ………………………..……………………….……………

41

4.3 Results and discussion ………………………………..………………….…………..

41

4.4 Conclusions …………………………..……………………………………….………...

50

5. Photoacoustic microscopy to image model hair fol licle ………………...……...

51

5.1 Introduction ……………………………..………………………………….……………

51

5.2 Materials and methods ……………………………..…………….…………………… 52

5.2.1 Materials ……………………………………..………………….…………………. 52

5.2.2 Equipment …………………………………………..……………….…………….. 53

5.2.3 Preparation PLGA NPs loading aminohexanamine stabilized magnetite …... 53

5.2.4 Preparation PLGA NPs loading oleic acid stabilized magnetite ……………... 54

5.2.5 Purification and charachterization of NPs ……………………………………… 54

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5.2.6 Determination of magnetite core ……………………………………...………… 55

5.2.7 Preparation of model hair shaft ……………………………...………………….. 55

5.2.8 Imaging of model hair shaft by CLSM and photoacoustic microscopy ………

56

5.3 Results and discussion ……………………………………..……….………………..

57

5.4 Conclusions ……………………………………………..………….…………………...

67

6. Summary ……………………………………………..………………….……………….

68

7. References ……………………………………………………………………………….

71

8. Abbreviations ……………………………………………………………………………

87

9. Curriculum Vitae ………………………………………..………….…………………...

89

10. Acknowledgements ……………………………………………………………………. 92

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SHORT SUMMARY

Caffeine in cosmetic products such as shampoo and lotion can stimulate hair growth.

Recently, it was shown that nanoparticulate formulations have better penetration into

hair follicles. Moreover such particulate formulations to stimulate hair growth are not

available commercially. Therefore in this study, two systems have been developed.

The first was a particulate formulation loading caffeine by utilizing biocompatible and

biodegradable polymers such as poly(lactic-co-glycolic acid) (PLGA) and chitosan.

The second was a model of hair shafts containing nanoparticles (NPs) to prove that

such model hair follicles can be visualized by photoacoustic microscopy. Results

revealed that there is an interaction between chitosan and caffeine suitable for drug

loading. Fourier Transform Infrared Spectroscopy (FTIR), Differential Scanning

Calorimetry (DSC) and solubilization showed that this interaction is based on

complex formation. NPs characterization showed that NPs have a spheric shape.

NPs from chitosan-PLGA showed good properties in terms of particle size and

distribution. Loading of caffeine was increased by using chitosan up to 19% EE.

Caffeine release from NPs is slower than from the pure complex allowing a longer

time frame for continious drug release. For imaging of NPs in model hair follicle

(voids in agarose gel), PLGA NPs loading magnetite were prepared. These NPs

could be successfully imaged by photoacoustic microscopy inside the model hair

follicle.

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KURZZUSAMMENFASSUNG

Koffeinhaltige Haarwaschmittel, können bekanntermaßen das Haarwachstum

stimulieren. Das Koffein wirkt dabei im Haarfolikel. Seit kurzem ist bekannt, dass

partikuläre Formulierungen topisch appliziert besser in Haarfollikel penetrieren als

Lösugen. Derzeit ist jedoch keine koffeinhaltige, partikuläre Darreichungsform

verfügbar. In der vorliegenden Studie wurden zwei unterschiedliche Systeme

entwickelt. Das Erste war eine partikuläre Formulierung mit Koffein. Hierzu wurden

biologisch abbaubare und kompatible Polymere wie PLGA und Chitosan verwendet.

Das zweite System sollte es ermöglichen die Partikel im Haarfollikel mittels

photoakustischer Mikroskopie zu visualisieren. Dafür wurde ein Modell-Haarschaft

entwickelt, sowie Partikeln, die einen guten Kontrast ermöglichen, hergestellt. Die

Interaktion zwischen Chitosan und Koffein wurde mittels Infrarot-Spektroskopie,

Kalorimetrie, sowie mittels Löslichkeitstest untersucht. Die Wechselwirkungen

zwischen Chitosan und Koffein beruhen auf Komplexbildungen und erlauben eine

Beladung der Partikel (19%EE). Die Partikel weisen eine geeignete Größe,

sphärische Formen und schmale Größenverteilungen auf. Freisetzungsversuche

zeigten eine verzögerte Freisetzung von Koffein aus NP im Vergleich zu reinen

Komplexen. Zur Visualisierung der Partikel in Modellhaarfollikeln (Hohlräume in

Agarosegel), wurden diese mit Magnetit-PLGA NP beladen. Die Partikel konnten

mittels photoakustischer Mikroskopie in den Modelfollikeln visualisiert werden.

I. Introduction

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I. INTRODUCTION

1.1 Hair loss problem in human life

Hair loss can happen to men and women. Patients are usually afraid of

experiencing it, because hair loss can lead to baldness [1]. The influencing factors

causing hair loss are shortly reviewed in terms of the conditions of the hair in the

follicles and the biosynthesis of hair which impact to the fragility of hair. Testosterone

and dihydrotestosterone are androgenic hormones which are involved on hair loss.

Overall, hair loss is influenced by internal factors which ranging from enzymes to

hormones and external factors [2-5].

1.1.1 Internal and external factors causing hair loss

Internal factors involve genetic reasons controlling the metabolism of

hormones and active substances relevant for hair growth. The gene Fgf5 is

responsible to encode the transcription factor for expression of fibroblast growth

factor 5 (FGF5). This growth factor plays an important role in the proliferation of hair

follicles. Three types common of baldness are due to genetic reasons: The first is

androgenic alopecia which can be seen anytime after puberty [6,7]. Usually the

problem will increase by increasing age. The second is telogen affluvium which is

often associated with genetics and hair loss occurs especially due to many hair

follicles suddenly stopping growing because of hormone imbalance (e.g., pregnancy)

[6,8,9]. Both these types of baldness happen in sequence until all hair is lost. The

third type is alopecia areata as shown in Fig.1 which indicates the spot or patchy

baldness [10,11].

Other internal factors were thyroid imbalance, allergic reactions and diabetes

mellitus. When thyroid imbalance occurred, high or low thyroid concentration can

interrupt the biosynthesis of testosterone and dihydrotestosterone that influences the

hair growth. Hair loss happened when testosterone is converted to

dihydrotestosterone [12]. Allergic reaction as the manifestation of hypersensitivity

reaction impacts on the activation of progressive fibrosis of the perifollicular sheet

occurs in lesions [13,14]. Diabetes mellitus can also cause hair loss. The growth of

hair is influenced by the concentration of glucose which can interact with keratin of

hair follicle which is known as glycosylation. Diabetes also influence the blood

I. Introduction

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circulation to hair follicles affecting the ability of the hair follicle to form metabolites

[15-18]. This also reduces the likelyhand of hair growth due to malnutrition [15-17].

Figure 1. Patchy hair loss “common type of alopecia” (image included with permission of volunteer)

External factors are the factors out of the human body such as substances

and environmental conditions which impact on the human body. External factors

involve psychic disorder, consumption of drugs such as chemotherapeutics and such

simple issues as hair styling. Psychic disorder because of stressful daily life can also

lead to several diseases including hair loss. Many works in modern life style lead to

less spare time for cooling down the metabolic reaction of the body. To support these

activities, the body also needs good nutrition for respective metabolic action. Lack of

protein and vitamin intake can cause hair loss. Therefore consuming sufficient

nutrition is necessary. Especially vitamins of vegetables and fruits play a role in

supporting the strength of hair root [19]. Consumption of anti cancer drugs such as

derivatives of cisplastin, aclarubicine and doxorubicine reduce the proliferation of hair

cells by being toxic especially to proliferating cells [20-22]. Hair styling is another

factor which impacts on the growth of hair especially when chemicals and mechanical

equipment are involved [23].

1.1.2 Prevention and treatment of hair loss

The natural way to prevent hair loss is to make it become healthy. Low hair

vitalization and dandruff formation can increase the probability of hair loss. All efforts

in order to maintain vitalization and to diminish dandruff formation are traditional way

that people applied since long time ago [24,25].

The treatment of hair loss is typically connected with an effort to reduce the

concentration of dihydrotestosterone [26]. On stem cells, there are receptors which

can bind testosterone. Afterwards, stem cells in the presence of testosterone [27]

induce the production of specific ligands interacting with natural killer cells (NK cells).

The ligands are proteins of stem cells also activating monocytes which are then

I. Introduction

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involved in hypersensitivity and cause inflammation as trigger for the degradation of

the root of hair follicle. As consequence, hair follicles can not grow [28]. The condition

when the hair follicle can not grow is known as miniaturization of the hair follicle. At

the basal layer of hair follicles the dermal papillae is located. It contains fibroblasts

which can regulate hair growth. The fibroblasts have androgen receptors. The larger

sensory nerve branches and the blood vessels that nourish the skin are also located

in the dermal papilae. This layer is an important part to inhibit the change of

testosterone to dihydrotestosteron by using active substances such as caffeine

[29,30] . Based on this information, the strategy to treat the hair loss is by inhibiting

the conversion of testosterone to dihydrotestosterone. For this purpose, caffeine and

also estradiol have been used [26,31]. Treatment of hair loss has also already been

conducted by using commercial products such as lotions, creams, hair sprays and

shampoos [32].

1.1.3 Caffeine as stimulating agent to stimulate hair growth

Rubiaceae plants are used in pharmaceutical application since long time ago

as well as in beverages. These plants have several bioactive compounds such as

caffeine which belongs to the alkaloid purine. As an alkaloid, Caffeine is obtained in

varying quantities in beans, leaves and fruits of plants such as coffee, tea, cacao,

and cola [33]. The presence of caffeine in those plants is to defend against

pathogens. People use the leaves of tea (Fig.2) for common consumption as well as

coffee, cola and cocoa bean. These plants are also used in commercial products.

These products are legally unregulated in nearly all jurisdictions even though

containing psychoactive substances [34,35].

Figure 2. Camellia sinensis is well known as tea plant. Image was taken in private garden in Bandung, Indonesia.

I. Introduction

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Caffeine consumption improves the physical endurance towards reduced drowsiness

and restoring alertness, cognitive function, particularly vigilance, mood and

perception of fatigue [25,33] .

Caffeine is a white powder with a hexagonal crystal structure of alkaloid

purine. Caffeine has a molecular weight of (Mw) = 194.19 Da and the monohydrate

has the Mw = 212.12 Da. The melting point of caffeine is 238°C. As can be concluded

from Mw which was schetched in Fig.3 the chemical name is 1H-purine-2,6-dione,

3,7-dihydro-1,3,7-trimethylpurine (C8H10N4O2) with C 49.48%, H 5.19%, N 28.85%, O

16.48%[36] .

Figure 3. Molecular structure of caffeine

As a central nervous system (CNS) stimulant, caffeine can increase brain

activity by blocking the receptor for neurotransmitters such as dopamine, serotonin,

acetylcholine, glutamate and γ-aminobutyric acid. Regarding hair growth, caffeine

does not act directly to the main pathway of hair synthesis. However, caffeine and

cAMP have a similar structure. Under physiologic condition, the concentration of

cAMP is decreased by dephosphorylation reaction. In the presence of caffeine,

dephosphorylation can be inhibited and therefore the concentration of cAMP is

increased [31,37].

1.2 Topical and follicular nanoscale drug delivery systems

The barrier function of human skin imposes physicochemical limitations to the

permeation of drugs that can cross this barrier. For a drug to be delivered passively

via the skin is difficult. An adequate lipophilicity is necessary to enhance the

permeation [38,39]. However, a strategy has been developed to direct delivery based

on particulate formulations to hair follicles [40]. This application has an advantage for

an implementation of topical and follicular drug delivery systems. The investigations

I. Introduction

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have been conducted to evaluate this new, innovative, and convenient dosage form

to target hair follicles [41].

1.2.1 Topical nanoscale drug delivery systems

Topical drug delivery systems are systems used to mainly apply drugs on the

skin to obtain localized effects at the site of application. Topical drug administration is

supported by the ease of administration and need to take into consideration the skin

structure as described in the following. Skin as the largest organ, has an area of 1.7

m2 and approximately 4 kg in weight or about 5.5% of the body mass [41,42].

Figure 4. A schematic sectional view of skin which involves stratum corneum (SC), viable epidermis and dermis (Image is adapted from [43]) Skin is known as the outer barrier between the body and the environment and

protects the body from external chemicals and pathogens. Skin is made up of three

cellular layers as is shown in Fig.4. Each of them has its own structure and function.

The outermost cellular layer of the skin is the epidermis which is composed of the

viable epidermis consisting of living cells and the non-viable stratum corneum (SC):

cornified cells forming a densely packed layer being the strongest barrier of the skin.

The dermis lies directly underneath the epidermis and consists of compact

connective tissue nerved with blood and lymph vessels. This formation supplies the

epidermis with nutrients and removes absorbed exogenous substances acting as

sink. The subcutaneous fatty tissue in dermis as well as other skin layers, consist of

loose connective tissues, and its dimensions vary greatly. Topical drug administration

Stratum Corneum

Viable epidermis

Dermis

I. Introduction

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gained attention as the skin offers an easy and hence convenient route having great

potentials to deliver drugs compared to other drug administration such oral, rectal

and parenteral [44].

Interestingly, for topical application, nanoparticles (NPs) are already used in

cosmetics [45]. Talking about terminology of nanoparticle, nano means small

originating from the Greek term for dwarf. Materials which have dimensions between

1 and 100 nm can possibly show unique properties enabling novel applications.

Nanotechnology is the creation or use of the nanometer-sized materials [46,47]. In

this thesis the usage of nano’ will be extended with respect to nanomedicines where

also sub-micron ranges are considered to be part of it.

Although NPs in cosmetics are commercially available, it is still under debate if

penetration into the skin is happening and to which extend. Nevertheless, several

results of investigation about penetration NPs across the skin were reviewed [48-50].

Several studies investigated the penetration of inorganic NPs such as gold NPs [51],

lipid-based NPs [52] such as liposomes [53], transferosomes [54], ethiosomes [55],

solid lipid nanoparticles (SLNs) [56], nanostructured lipid carriers and surfactant

based systems such as nanosomes [57], micelles [58], and nanoemulsion [59].

Based on the investigations regarding the penetration of NPs into the skin, 6

nm AuNPs showed much higher extent than 15 nm AuNPs. Furthermore, it indicated

a minimal effect of the vehicle on particle penetration [51,60,61]. The investigation of

topical application of Fluorescein-PLGA NPs with size 320 nm reported that particles

were only distributed on the surface of the skin. The measurement was conducted by

CLSM and revealed that particles located near to the lipid layer around the

corneocytes [62]. In conclusion, only very small particles seem to be able to

successfully penetrate into the deeper skin layers. The determination of the amount

of NPs in the SC could be done by tape-stripping [62,63]. Furthermore, the

experimental set up for a penetration experiment for NPs were also studied. For

instance, exposure times of at least more than 6 hours were recommended for future

studies on skin penetration of NPs. These obtained informations are very important

for the basic understanding of the interaction of NPs with the skin barrier [51,64]. This

would be of use for future pharmaceutical and clinical applications, e.g. designing

optimal topical and transdermal delivery systems. Additionally, some drugs have

been evaluated to be delivered into the skin; for instance the evaluation of anti aging,

vaccine and anti malignant melanoma [65-67].

I. Introduction

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1.2.2 Penetration pathways across the skin

Substances delivered into the skin involve three major pathways as shown in

Fig.5. The possible routes are transcellular, intercellular and transfollicular pathway.

The transcellular pathway is considered to be of minor importance for dermal

absorption due to low permeability of certain substances through the corneocytes

[68]. In this transport pathway, substances have to partition from hydrophilic

corneocytes to the lipid layers of the SC repeatedly resulting in a very slow process

[69]. The intercellular pathway is considered as the predominant pathway for most

substances. In this case, substances diffuse within the continuous intercellular lipid

domains of SC [70,71]. In contrast to the corneocyte structure which is compact, the

lipid domain pathway can absorb the substances faster than corneocytes and also a

higher amount of the substances can be absorbed. Furthermore, this pathway can be

an alternative especially in the presence of penetration enhancers [51]. It is known

that the permeability of corneocytes increases due to alteration of keratin structure.

However, nowadays the transcellular pathway has been investigated and may be

more relevant to evaluate new formulations for dermal therapy [72].

Figure 5. Scheme of penetration pathways (Image is adapted from [73])

As obvious from Fig.5 the third pathway is the skin appendages offering a

direct passage into deeper skin layers. A high density of blood vessels around the

hair root and the absence of the SC make this an intriguing pathway especially for

Blood vessels

Dermis

Epidermis

Stratum Corneum

Drug

Transcellular Intercellular Transfollicular

I. Introduction

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the hair follicles also showing the possibility to be permeated by drugs [74-77]. Some

research was focused on the evaluation transfollicular drug delivery [78]. More

detailed explanations about transfollicular pathway will be given in the next chapter

1.2.3.

1.2.3 Transfollicular pathway for drug delivery systems

Transfollicular pathway can deliver drug substances and particles into hair

follicles. A respective image of a hair follicle is shown in Fig.6. Hair follicles are

embedded in the epidermis extending deep into the dermis which provides a much

greater actual area for potential absorption below the skin surface.

Figure 6. Light microscopy image of hair follicle (image was depicted with help of Christiane Mathes at Biopharmaceutics and Pharmaceutical Technology, Saarland University). The structure of the hair follicles is described as a complex structure which is formed

by three common parts: dermal papilae, bulge and sebaceous, and also the

infundibulum [79]. Each part represents the distinct program of differentiation for

follicle morphogenesis during the embryonic cycle. The dermal papilae in basal layer

is the most important layer. This layer is covered by the follicular ephithelium. The

dermal papillae contains specialized cells, so-called fibroblasts, that regulate hair

growth. Above this layer, are located the bulge area and the sebaceous glands [80] .

The next part is the infundibulum. From recent evidence about regulation of hair

growth is known that the infundibulum area also has a function to regulate follicular

growth and differentiation [81]. The location of bulge, sebaceous gland and

infundibulum are schematically shown in Fig.7.

Cosmetic products have been applied on the surface of skin but

pharmaceutical dosage forms are facing the problem to deliver drug into the skin. To

bring the drug across the stratum corneum is not an easy task. However, hair follicles

Hair Follicle

I. Introduction

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give a significant contribution to the penetration which is known as transfollicular

pathway [82,83]. In the past decade there was not much attention to this pathway

[84]. A study about non particulate formulation was done to investigate its ability to

penetrate hair follicles. The substance chosen for this study was estradiol. It is a

poorly soluble, neutral compound with log octanol-water partition coefficient of logP =

2.29 and a water solubility of 0.0003%. In 2002 the sandwich model with SC

membranes for a Franz diffusion cell experiment was described. By using this SC

sandwich, putting two sheets of SC on top of each other, there is only a negligible

chance for a direct connection of two openings (hair follicles) across both membranes

because of the random distribution of hair follicles on the skin surface. The result

showed that the permeation through the sandwich was much reduced rather than

that of a single skin membrane (SC) [85].

Figure 7. Scheme of sebaceous gland, bulge, hair matrix and infundibulum of hair follicle (Image is adapted from [86])

The group of Jürgen Lademann at the Centre for Experimental and Applied

Cutaneous Physiology, Charite University of Medicine Berlin, Germany, investigated

the delivery of cucurmine as substance for topical application. The presence of

cucurmine was determined by using confocal laser scanning microscopy (CLSM) of

skin biopsies. It could be shown that the substance could penetrate by using follicular

pathway [83].

Bulge

Sebaceous gland

Infundibulum

Dermal papillae

I. Introduction

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Furthermore, the transfolicular pathway was found to be accessible for sub-

micron particles. Hence these systems can be used as drug delivery systems to

specifically target the hair follicles. This specificity might turn the nano- and sub-

micron carriers into important drug delivery system for the skin. A study about the

penetration of TiO2 microparticles contained in sunscreens was conducted to

determine their ability to penetrate hair follicles by using tape-stripping as well as by

using X-ray fluorescence microscopy. It could be shown that these particles

penetrated into the hair follicles [87] . Further investigations have even demonstrated

that NPs have a better penetration than non-particulate formulations[80] . Regarding

the particles size, microparticles of size 3–6 µm showed a tendency to aggregate in

the hair follicles [88,89] . Particles in the size of 750 nm showed a homogenous

distribution in the hair follicles [88]. Lademann reported in 2009 that particles sized

300–600 nm penetrate efficiently into hair follicles than larger particles [90]. This size

range corresponds to the approximate size of the hair cuticula which is 530 nm for

human hair and 320 nm for pig hair (which is assumed to play an essential role in the

permeability process by a pumping process due to the movement of the hair) [90].

Also particles of ~100 nm in size were found to be able to penetrate into the

hair follicles after sunscreen application [86]. Even smaller particles with sizes as

small as 40 nm were found to penetrated deeper into the follicle and could also reach

the follicular epithelium [91]. These small particles allow also to penetrate in the

sourrounding tissue. According to the definition of NPs as particles being around 1–

100 nm it can be concluded that NPs are better than microparticles to address

cellular internalizsation around the hair root for instance in langerhans cells for

possible vaccination [92].

The underlying mechanism for the superior particle penetration was explained

to be due to the hair working as a geared pump in the hair follicle area. Hence the

movement of the hair drives the particles deep along the hair into the hair follicle. On

the other hand, the investigations conducted by Lademan in 2006 and Otberg in

2007 revealed that hair follicles also represent an efficient storage for long term

which keep those particles in reservoir [62,80].

I. Introduction

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1.3 Particulate formulation as drug delivery system s

Nanotechnology has found applications in nearly all fields leading to significant

technological advantages that can be applied such as in the pharmaceutical field for

preparing and characterizing nanoparticulate formulations. Particulate formulations

are stabile products and they have large potential to target and control the release of

encapsulated drugs for instance due to different ways of interaction. Many efforts are

focused on the development of these particulate formulations [93].

1.3.1 Current status of polymeric nanoparticles

Polymeric nanoparticles (NPs) are NPs which were prepared by using

polymers. For pharmaceutical application, biodegradable and biocompatible

polymers are preferred. The advantages to use such polymers are because of that

the degradation of these polymers to biologically acceptable molecules that are

metabolized and removed from the body using normal metabolic pathways. These

biodegradable polymers also are well known as safe and bio-tolerant because their

products of degradation and by-products fulfill the request for little or no adverse

reactions within the physiology of the human body. The success of loading polymeric

NPs with drugs shows that nanotechnology can provide carrier systems which can

and will be used for many exciting products potentially overcoming many hurdles in

formulation technology [94].

The drug is entrapped, encapsulated or attached to the NPs. Depending on

the method of NP preparation, nanospheres or nanocapsules can be obtained [95].

In recent years, biodegradable polymeric NPs based on natural polymers such as

chitosan, gelatin, alginate or synthetic polymers such as poly(lactic-co-glycolic acid)

(PLGA), poly (anhydrides), poly (caprolactone), poly (ortho esters), and poly (amino

acids) have attracted considerable attention as potential drug delivery systems in

view of their applications in drug targeting especially to particular organs/tissues

[96,97]. Among these polymers, PLGA is most often used and was already used in

implant medical products [98].

Some researchers used PLGA in laboratory scale to form NPs and it is due to

its benign nature a promising material to be used in future pharmaceutical products.

As can be seen from the structure, PLGA is condensed by a lactic acid block and

glycolic acid block by ester bonds forming a block copolymer (Fig.8).

I. Introduction

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Figure 8. Molecular structure of poly(lactic-co-glycolic acid) (PLGA)

The molecular weight of PLGA ranges from 5,000 to 50,000 depending on the

composition of lactic and glycolic acid blocks. The higher the number of lactic and

glycolic acid blocks or the respective lactide block, the bigger is the polymer. PLGA is

a semipolar polymer which dissolves in semipolar solvents such as ethyl acetate,

acetone, and dichloromethane. For the preparation of nanoparticles [96,99], PLGA is

often disolved in ethyl acetate as an organic phase and the stabilizer is added into

the aqueous phase [96].

For investigation of drug delivery, particles based on poly lactic acid

derivatives such as PLGA are used. These particles could be appreciable also for the

delivery both of hydrophilic and hydrophobic drugs. However, in the past it was

observed that hydrophobic drug can be loaded into the polymeric NPs very well.

However, for hydrophilic drugs, the situation is more complicated and a double

emulsion method is preferred to incorporate hydrophilic compartments containing the

drug in the particles [100]. Another method such as nanoprecipitation is also used for

hydrophilic drugs [101,102]. As possible hydrophilic pharmaceutical active agents,

therapeutic proteins and vaccines were used. Until today, there are still limited data

concerning PLGA NPs containing hydrophilic substances [103]. Looking in the

literature, it turns out that PLGA is often applied as a drug carrier system to study the

delivery and targeting of colon cancer drugs. Nevertheless, its biodegradability

makes it also a useful material for other application routes. The PLGA NP

formulations prepared are charachterized in terms of size, dispersity index, zeta

potential, and release profile [104]. The adjustment of these physical properties is the

key for targeting as they correspond to the interaction with the target location of the

NPs. The loaded hydrophilic drugs which were studied ranged from antibiotics, anti

cancer drug, anti inflammation drugs, and therapeutic proteins including vaccines

[48,105,106].

Besides PLGA, chitosan, gelatin and alginate are also relevant and important

substances to form polymeric NPs. These polymers were grouped as natural

I. Introduction

- 18 -

polymers. In contrast to the polymeric NPs based on synthetic polymers the natural

polymers show a broad size distribution. Hence for basic understanding and

investigation they are less suited and particles from synthetic materials such as

PLGA are often preferred due to their better size distribution [107,108].

1.3.2 Preparation of polymeric nanoparticles

The PLGA NPs are appropriate for drug delivery systems and have been

applied because of its biodegradability ensuring that the carrier itself and its products

do not disturb the physiologic conditions of the human body. This pharmaceutical

important ability resulted already in a pharmaceutical product containing PLGA,

which is being approved by national food and drug administration (FDA) [98] hence

underlining the materials’ potential. Therefore, these NPs are used as vehicles for the

targeted and controlled delivery of drugs. Investigations showed successfully that

PLGA NPs are appropriate for various routes drug administration [96]. To form these

NPs methods know from nanotechnology are used. This technology is appropriate to

form NPs and also showed the ability to load during the process pharmaceutical

substances in the particles. NPs based on PLGA of different physical characteristics

such as size, distribution of particles, morphology and zeta potential can be

synthesized by controlling the specific parameters of the synthesis.

To load hydrophilic drugs is more difficult than hydrophobic drugs due to

miscibility problems between the drug and the particles’ material. Nevertheless,

Barichelo et al. in 1999 intended to evaluate the loading of hydrophilic drugs (insulin

and valproic acid) in NPs which were formed by nanoprecipitation [102]. Another

hydrophilic drug such as procaine hydrochloride [103] was shown to be successfully

loaded into NPs.

It is known that the synthesis on NPs by available methods such as emulsion

solvent diffusion could be used as common method for preparation of NPs based on

PLGA. As shown in Fig.9 two phases -organic and aqueous- phase (which are

partially miscible) are involved. Typically, the polymers are soluble in organic solvent

and the surfactant as stabilizer is soluble in water as the aqueous phase. The organic

phase is dropped slowly into the aqueous phase under stirring condition. A pre-

emulsion is achieved after continuous stirring for 1 h. The size reduction of the

preemulsion the so-called nanoemulsification is conducted by transferring energy into

the system using mechanical processes such as ultraturrax (rotor-stator principle) or

I. Introduction

- 19 -

ultrasonifier from 1 minute up to 5 minutes depending on the expected size of the

particles (Fig.9). Dilution with water allows the organic solvent to diffuse into and mix

with water. As a result a solid particle from polymer without solvent is formed. After

that overnight evaporation with low vapor pressure is needed to remove the organic

solvent form the surrounding.

The double emulsion method is a common method to address the solubility

issues between polymer and drug and hence to encapsulate hydrophilic drugs.

Similar to the single emulsion solvent diffusion, but in this method two emulsification

steps are involved by using two kinds of stabilizers. Here the inner, first emulsion

contains the drug in an aqueous environment surrounded by polymeric material

which is then incorporated into another, larger water droplet. The dilution with water

is also required to extract the organic solvent overnight.

Another method to prepare polymeric NPs is nanoprecipitation which was also

applied for hydrophilic drugs such as therapeutic proteins [102,109]. Overall the

method is similar to the solvent diffisuoin method, but for this method miscible

solvents are needed instead of partially soluble solvents. The polymer is dissolved in

the organic solvent such as dimethyl sulfoxide (DMSO), ethanol or aceton. Stabilizer

is dissolved in aqueous phase.

The respective drug is placed in the solvent where it dissolves. When mixing

the organic phase containing the polymer with the stabilizer-containing aqueous

phase, the miscibility of the solvents leads to an immediate precipitation of the

polymer. As the polymer-solvent diffuses away, the polymer collapsed and

precipitates in the nano size because it is not soluble in water [100,102,103].

I. Introduction

- 20 -

Figure 9. Preparation of NPs by using emulsion solvent diffusion method with ultrasonifier. Polyvinyl

alcohol (PVA) is a stabilizer and the method can be expanded to also load magnetite particles (MT).

1.3.3 Chitosan and chitosan-coated PLGA NPs

Chitosan-coated PLGA NPs are formed by using chitosan and PLGA. The

coating involves positive charge of chitosan and negative charge of PLGA to mediate

the interactions [110,111]. Chitosan is a hydrolized extract of chitin from crustaceans’

hardshell, such as shrimp, crabs, insects, and also mushrooms. To obtain chitosan

the main processes which are involved are the following: first, the skin of shrimp or

crab was washed to be deproteinated. Then the sample is washed with acid solution

to remove the lime as demineralization process. After demineralization, the acetyl

groups of chitin were cleaved leading to chitosan. The ratio of deacetylation

corresponds to the positive charge coming from primary amine groups of chitosan

[112].

Chitosan is poly-D-glucosamine which is composed by more than 5000 units

of monomers (glucosamine and acethylglucosamine) with a molecular weight up to

500 kDa (Fig.10). The monomer number from which chitosan is composed is not less

than 16 [107]. For biomedical applications, chitosan is already used as wound

dressing to stop bleeding and is in addition applied due to its antibacterial properties.

PLGA

PVA

Stirring 1 h

Homogenizer

Dilution and Evaporation

Small drops Organic Aqueous 00

Peristaltic Pump

Ice bath

I. Introduction

- 21 -

Chitosan (chit) is generally soluble in acidic solution due to the protonation of

the amine groups. However chitosan chloride usually can also be dissolved easily in

water of neutral pH. Therefore, for the preparation of chitosan-coated PLGA NPs,

chitosan was added into the aqueous phase and with a simple one step emulsion

solvent diffusion, chit-PLGA NPs can be prepared [113]. These chit-PLGA NPs have

also been used for investigation regarding transfection of antisense oligonucleotides

and gene delivery due to their positive surface charge [110,114]. Besides for gene

therapy, chitosan NPs have shown a good performance to load other drug for

example rifampicin as antituberculosis agent [115].

1.3.4 Magnetite loaded polymeric NPs

The importance of nanotechnology has offered many opportunities in various

research fields. Based on imaging and therapy, particularly inorganic NPs have

received great attention because of their outstanding properties. Metal NPs have

many advantages over small conventional molecules that include high molar

extinction coefficient, high resistance to photo-degradation, size/shape dependent

and tunable absorbance/scattering properties, which can be useful for imaging and

therapeutic approaches. Especially the size/shape dependent and tunable

absorbance /scattering properties can enable on-demand design for imaging or

characterization purposes of many inorganic NPs such as magnetite [116].

Magnetite is iron oxide (Fe3O4) with superparamagnetic properties which

render interesting properties as important molecule than can absorb the near infra

red light and could be attracted with magnetic field. Paramagnetism is a form of

magnetism where certain materials are attracted by an externally applied magnetic

field. Paramagnetic properties are due to the presence of some unpaired electrons

and form the realignment of the electron orientation caused by the external magnetic

field. On the other hand, superparamagnetism is a form of magnetism which appears

Figure. 10. Molecular structure of chitosan with (a) deacetylated and (b) acetylated monomer

(a) (b)

I. Introduction

- 22 -

in small nanoparticles for instance magnetite particles. Their size is small enough that

only one magnetic domain exists per particles. Therefore the magnetization can

randomly flip direction at room temperature due to the thermal energy. When an

external magnetic field is applied, their magnetic moments are aligned along the

applied field. The interest in magnetic materials is due to their function to develop and

serve in modern technology [99,116]. Magnetite nanoparticles (MNPs) which possess

paramagnetic properties were already evaluated in the clinic as contrast agent [117]

or as therapeutic option in glioblastoma [118,119] . Furthermore, the ability to direct

and hence target these particles by local magnetic fields offers further potential

therapeutic applications [120,121].

Magnetite was also incorporated in polymer particles. In this case, the

magnetic moment of each magnetite will be able to rotate randomly in reference to

the orientation of the MNP. The important property for biomedical application was the

lack of magnetization after the colloid got stable to avoid the agglomeration [122].

The investigation which were conducted by Maity, 2007 and Dresco, 1999

presented the best compromise among appropriate magnetic properties such as

saturation magnetization, stability under oxidizing conditions and safety for biological

application [123,124]. The US Food and Drug Administration (FDA) has already

approved the medical product such as Feridex® and Resovist® containing magnetic

NPs which was formed by a mixture of Fe2+ and Fe3+ [125].

II. Aim of Dissertation and Experimental Design

- 23 -


The problem of hair loss resulting in baldness is a serious health problem. So

far there are cosmetic products as non particulate formulations such as shampoo and

lotion containing caffeine to stimulate hair growth. In addition, cleansing creams and

lotions containing chitosan intended to clean the skin are also available on the

market.

Even though particulate formulations were shown to penetrate better into the

hair shafts (the target for caffeine delivery) no particulate caffeine formulation was

described so far for such a purpose. As consequence, no cosmetic product is based

on particles for caffeine delivery. Therefore, this research was intended to develop a

particulate formulation using the natural polymer chitosan and the synthetic polymer

PLGA to form chitosan-coated PLGA NPs as a carrier system loading caffeine.

Hair follicles as potential target for those carrier systems require also the

imaging of particles which penetrate into the hair follicles. Therefore, the formulation

of nanoparticles which can be imaged in respective structures would be necessary.

To address the problems with respect to the size of the hair follicles for imaging, a

new approach based on photoacoustic microscopy was aimed. The follicular imaging

with photoacoustic microscopy needs the right marker for particle visualization and

therefore a drug carrier system containing magnetite was the next goal.

Before going to the rather complex in vivo systems (animal or human skin), the

ability to image the particles at all in hair follicles was in focus. Therefore a system

was developed to mimic hair follicles and to image the particles in this structure ex

vivo.

Besides the overall direction of the thesis a focus on the experimental

approach of the different chapters is described below :

As loading of the nanoparticulate formulation was tried to be accomplished based on

specific interactions between the drug and the carrier. This study investigates the


- 24 -

interaction between chitosan and caffeine by using FTIR and DSC measurement in

combination with a solubilization study.

√ It was aimed to see whether the interaction pattern involved covalent bonds or

rather a complex formation.

√ Once known the interaction pattern, the following work was aimed to know the

binding capacity of chitosan which interacted with caffeine by using

solubilization study.

After identifying the basic interaction parameters preparation and

characterization of chitosan-PLGA NPs, it was in focus to find a good formulation.

√ The most common steps to make NPs are intended to be loading hydrophilic drug

such as caffeine using emulsion solvent diffusion. As an organic phase has

used PLGA which was dissolved in ethyl acetate and aqueous phase was

chitosan and PVA solution. For control, double emulsion technique as a

common method to encapsulate hydrophilic drug was applied.

√ This further study also intended to characterize NPs using zeta sizer, SEM and

AFM.

√ The amount of caffeine in NPs was determined indirectly from the supernatant of

suspension and directly by dissolving NPs. The release profile of caffeine from

chitosan-PLGA NPs across a membrane was determined and compared to the

complex alone and caffeine diffusion across a membrane.

As conventional technique, such as light microscopy requires sectioning of the

sample. Another imaging approach such as photoacoustic microscopy was in focus.

With this approach, a material which can enhance the contrast such as magnetite is

needed. Therefore the third part of the thesis was thought to develop a particulate

formulation containing magnetite particles for improved contrast. This formulation

would be applied to image the model hair follicle. As mentioned above, this model is

necessary to establish rather than working with the complex systems of in vivo

measurements. The workpackages for this part were:

√ Preparation of PLGA NPs loading magnetite using two kinds of magnetite. First one

was magnetite which stabilized with aminohexanamine and the second one was

magnetite was stabilized with oleic acid. Single emulsion solvent disffusion


- 25 -

method was applied to the preparation of NPs. For the comparison the

nanoprecipitation method was used.

√ Characterization of PLGA NPs loading magnetite in terms of size, dispersity index

of particles, zeta potential, and morphology. The presence and successful

loading of the core of magnetite was determined by AFM and TEM.

√ Development of the model hair shaft and imaging by using photoacoustic

microscopy. Model of hair shaft was based on agarose. The idea was to create

voids in the size of human hair follicles. The model hair shaft should be filled

with suspension of PLGA NPs loading magnetite for imaging. The imaging

measurement was conducted by using photoacoustic microscopy in cooperation

with IBMT, St. Ingbert.

III. Interaction of Chitosan and Caffeine

- 26 -


3.1 Introduction

Several efforts were addressed to the development of cosmetic formulations

especially when using chitosan and caffeine [126,127]. Chitosan which is

commercially available in formulations is of high-grade quality made from shrimp

shell by deacetylation (75-90%). Furthermore due to the positive charge, chitosan

can interact with DNA and therapeutic proteins to avoid degradation and facilitate

cellular uptake [128-130]. Chitosan was also used successfully to form particles

containing theophyline derived from xanthine (alkaloid) as a lead structure [131,132].

However, there were no data regarding the interaction of chitosan and caffeine.

Based on the molecular structure of chitosan and caffeine, they both have

several polar functional groups which lead to hydrophilic properties. Usually polar

interaction of the drugs could be categorized into dipole-dipole (Keesom) and dipole-

induced dipole interaction (Debye) which involve dipole orientation of molecules with

polarizable dipole moments. Depending on the structure of the molecule, both types

of interactions can take place in the underlying van-der-Waals (vdW) forces

[133,134].

To study the interaction between chitosan and caffeine, the information about

their polarity is necessary. Their polarity refers to a separation of electric charges

leading to a molecule or its chemical groups having an electric dipole or multipole

moment. Polar molecules interact through dipole-dipole intermolecular forces and

hydrogen bonds and molecule polarity depents on the difference in electronegativity

between atoms in a compound and the asymmetry of the compound’s structure.

Furthermore, polarity of the molecules results in a number of physical properties

including surface tension, solubility, boiling and melting points [135,136]. This polarity

usually corresponds to hydrogen bonds forming between the hydrogen atom

attached to an electronegative atom such as oxygen in the carbonyl functional group

(C=O) of caffeine, and nitrogen in the amine group N-H2 of chitosan. Based on these

theories we assumed that chitosan and caffeine can perform this kind of interaction to

associate with eachother.


- 27 -

3. 2 Materials and Methods

3.2.1. Materials

Chitosan with 75% deacetylation and a molecular weight Mw < 150,000 Da

was obtained from Novamatrix, Norway (Protasan UPCL 113). Centrisart with a

molecular weight cut off (MWCO) of 20,000 Da, Vivaspin 20 which a MWCO of

300,000 and 100,000 Da were obtained from Sartorius, Göttingen, Germany.

Caffeine anhydrous was obtained from Sigma Aldrich, St Louis, USA. All other

solvents and chemicals were commercially available, from the highest grade and

used as obtained.

3.2.2. Equipment

UV-Vis Spectrophotometer, Lambda 35 and FTIR Spectrophotometer,

Spectrum 400 series, PerkinElmer LAS Rodgau, Germany were used. Differential

Scanning Calorimetry (DSC) was performed by using DSC-Q100 from TA Instrument,

Germany. Sample holder for DSC from Hermetic Lid, Germany, Sonicator, Bandelin

from Sonorex, Germany, Vortex Genie from Scientific Industries, Germany,

Centrifuge, Rotina 420R from Hettich Zentrifugen, Germany, and pH meter from

Schott, Germany was used for the different investigations.

3.2.3. FTIR measurement

To perform Fourier transform infrared spectroscopy (FTIR) measurements,

physical mixture of chitosan and caffeine powder was used. For the comparison, the

samples was also prepared from solution by dissolving Protasan UPCL 113 at pH 9

to obtain 0.3% chitosan solution with 10 mg of caffeine. While stirring, the interaction

could take place for 30 minutes at room temperature followed by lyophilization. A

total of 3 mg of this mixture was characterized by FTIR Spectrophotometry. Pure

chitosan and caffeine anhydrate were also used as references.

3.2.4. DSC measurements

DSC measurements were taken under nitrogen flow of 50 -100 mL.min-1 using

a sample mass of 4 mg and heating rates of 10°C min-1. The samples were placed

into covered aluminum holders with a central pinhole. An empty sample holder was

used as reference and the runs were performed by heating the samples from 25 up


- 28 -

to 400°C. The samples were the physical mixture of chitosan and caffeine, chitosan

(Protasan UPCL 113) and caffeine were used for reference.

3.2.5 Determination of chitosan and caffeine by UV spectroscopy

Stock solution of chitosan chloride was obtained by dissolving 5 mg Protasan

in 2 mL demineralized water. After that, into the chitosan solution, hydrochloride acid

0.1 M was added to adjust the concentration of chitosan to 0.25 mg/mL. From this

solution, the determination of λmax was performed by UV spectrophotometer. The

measurement was performed in quartz cuvettes between 200 and 300 nm using an

absorption peak of chitosan [137]. For calibration, serial concentrations of chitosan in

hydrochloride acid 0.1 M was used to obtain the concentrations of 0, 0.006, 0.125,

0.250, 0.350, 0.500, 0.650 and 0.750 mg/mL. For the solubilisation study, the

permeation of chitosan through different filters was tested (Vivaspin 20 with MWCO

300,000 and 100,000 Da and Centrisart with MWCO 20,000 Da) and the

concentration of chitosan passed through the membrane was calculated. For the

preparation of caffeine solution, 5 mg caffeine was dissolve in demineralized water as

stock solution. For calibration concentrations of caffeine of 0.0015, 0.003, 0.004,

0.006, 0.008, 0.012 and 0.024 mg/mL were used

3.2.6 Solubilization study of caffeine in the presence of chitosan

First of all, caffeine in high amount was added to an aqueous phase at 25°C to

determine the saturation concentration (cs) keeping the temperature constant. For

solubilization studies, the amount of caffeine was always kept higher than cs.

Figure 11: Scheme of the setup which was used for the determination of cs of caffeine using Centrisart 20 kDa to measure the solubility of caffeine.

Membrane 20 kDa

Chitosan with soluble caffeine Soluble caffeine Insoluble caffeine


- 29 -

Determination of the dissolved amount of caffeine was conducted by adding different

amounts of chitosan to the inner chamber of the Centrisart (Fig.11). A total of 60 mg

of caffeine were suspended in 1.5 mL of demineralized water. The sample was

transferred to the outer tube of the Centrisart. Then 0.5 mL of chitosan with several

concentrations 0.075, 0.150 and 0.300 mg/mL were added separately to the inner

tube with membrane MWCO 20,000 Da as shown in Fig.11 and followed by

centrifugation with 1000 x g for 90 minutes. Hereafter, the amount of caffeine in the

supernatant was determined by UV spectrophotometer.

3.3. Results and Discussion

3.3.1. Physical interaction between chitosan and caffeine

Protasan UPCL 113 used in this experiment, is commercially available, has an

average degree of deacetylation of more than 75% and its molecular weight

distribution ranges from 10,000 to 400,000 Da. This type of chitosan chloride is

soluble in water also in pH range 5 to 9. At pH 11 Protasan precipitates. In the pH

range where Protasan is soluble its amine (-NH2) functional group can possibly

perform an interaction with the carbonyl group (C=O) present in caffeine.

The determination of an interaction between chitosan and caffeine is the first

indication which is a necessary for the loading of caffeine into chitosan-PLGA NPs.

Therefore, the interaction between chitosan and caffeine was addressed looking at

IR spectra and the glass transition temperature.

First of all, the samples were prepared and the interaction patterns were

analyzed by using a procedure to determine caffeine [138]. As shown in Fig.12, the

FTIR spectra display the typical peaks from the functional groups. The stretching

bond of carbonyl is typically found in the range between 1447-1705 cm-1 [139]. For

caffeine, the stretching bond of carbonyl was found at 1646 cm-1 right in the range as

expected. The stretching bond of methylamine is typically found in the range between

992-1260 cm-1 [140]. For chitosan, stretching bond of methylamine was found at

1020 cm-1 also right in the range as expected. Both wavenumbers did not change

their position for the mixture. The complex formation is supported by the change of

intensity of the peaks in the spectrum allowing to conclude that the interaction

involves hydrogen bonds between the amine groups of chitosan and the carbonyl

groups of caffeine. Furthermore, it can be concluded, that the interaction does not


- 30 -

involved formation of covalent bonds as there was no shift in the wavenumber (cm1)

observed (Fig.12). Possible covalent bonds would occur between amine and carboxyl

groups (HO-C=O) in the presence of high temperature. As caffeine only has a

carbonyl group and the interaction took place at room temperature (~23°C) the

formation of a covalent bond was not likely. The wavenumber above 2750 cm-1

represents the stretching of –OH and C-C of chitosan. Blue lines are the complex of

chitosan and caffeine and the stretching –OH and C-C were revealed as shown in

Fig.12.

In Fig.13 the DSC measurements are displayed. It can be seen that water evaporates

at the peak below 100°C in the chitosan spectrum (grey line). Typically this water is

associated with hydrophilic groups in the amorphous chitosan. Caffeine as anhydrous

base did not show that peak (black line) as expected. Endothermic phase transition

from solid to liquid took place below 250°C for chitosan and caffeine. The

disappearance of the endothermic peak of caffeine at 310°C indicated that caffeine

dissolved in the molten chitosan which is an indication that chitosan and caffeine

form a complex.

40

50

60

70

80

90

100

110

500100015002000250030003500

w avenum ber (cm -1)

mixture of chitosanand caffeine

chitosan

caffeine

freeze dried ofchitosan andcaffeine

wavenumber (cm-1)

Figure 12 : FTIR spectra of chitosan, caffeine and chitosan-caffeine as physical mixture and prepared from solution. The respective wavenumbers 1020 and 1646 cm-1 are highlighted by thed dashed line indicating the strechting of the amine group and the carbonyl group, respectively.


- 31 -

Figure 13. DSC measurements of pure chitosan, caffeine and both together to identify a possible physical interaction between caffeine and chitosan

3.3.2 Solubilization study of caffeine by using chitosan

This step was performed to support the information that was already obtained

regarding the physical interaction that was explained above. At first, the change of

dispersion turbidity containing saturated caffeine was performed. Fig.14 shows that

(Fig.14A), the saturated solution containing insoluble caffeine results in a turbid

solution. Afterward, chitosan was added to this solution of caffeine resulting in clear

solution (Fig.14B).

Chitosan

Caffeine

Chitosan-Caffeine

Figure 14. Saturated caffeine become clear solution because of solubilization by adding chitosan. A. Saturated solution containing caffeine

B. After adding chitosan

A B

4

5

6

7

8

9

10

11

12

13

0 50 100 150 200 250 300 350 400 450

Temperature ( 0C)

Heat Flow (mW)

chitosan

caffeine

chitosan-caffeine


- 32 -

This reduction in turbidity is caused by additional dissolved, suspended caffeine by

the presence of chitosan. Since the free caffeine interacts with chitosan, forming

complexes, the non-dissolved, solid caffeine can dissolve to reach the saturation

concentration.

Besides this first indication regarding turbidity, the capability of chitosan to

solubilize caffeine was investigated. To test the solubilization capability of chitosan

for caffeine, a filter setup was used. In this setup caffeine and chitosan were in two

different compartments separated by a membrane. For the compartments Vivaspin

system or Centrisart system was tested. Vivaspin has one tube and in that tube

located the membrane with certain MWCO (300,000 and 100,000 Da). Protasan

UPCL 113 has molecular weight <150,000 Da. The amount of chitosan passed

through the membrane can influence the amount of caffeine which was solubilized in

this experiment. Therefore, before going to the solubilization study, the determination

of chitosan passing the membrane is necessary. Chitosan was measured using the

absorption maximum at 207 nm in acidic solution of pH 1 (0.1 M HCl) [137]. At high

wavelength close to 300 nm there was no absorption as shown in Fig.15. Therefore,

the peak at 207 nm works well for the determination and quantification of chitosan as

can be seen from the Fig.15.

Figure 15. Absorption maximum of chitosan between 200 and 300 nm and the respective calibration curve of chitosan based on this absorption.

Based on result in Fig.16, Vivaspin is not suitable for solubilization study

because chitosan can pass through the membrane. On the other hand, Centrisart

presents small MWCO to hold chitosan (Fig.16). With centrisart (20,000 Da) system,

chitosan has not passed through the membrane.

-0.05

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

195 215 235 255 275 295

Wave lenght (nm )

A

207

y = 1.9281x + 0.0147

R2 = 0.9977

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8

c o n c e n t r a t i o n o f c h i t o sa n ( m g / m L )


- 33 -

Figure 16. Amount of chitosan that could pass through the filter of Vivaspin and Centrisart with different MWCO.

Centrisart (20,000 Da) was investigated furthermore to perform solubilization study

(hence allowed to determine the caffeine concentration with interference from the

polymer). Subsequently, determination of chitosan and caffeine can be determined

by using UV spectrophotometry.

In Fig.17 we can see the peak of caffeine which also could be determined by

using UV spectrophotometry. Caffeine has the maximum absorption at 272 nm in

demineralized water. Based on the two spectra of chitosan and caffeine, it was

concluded that chitosan did not interfere with the caffeine determination because

chitosan does not absorb at 272 nm (Fig.15).

Figure 17. Local absorption maximum of caffeine at λ= 272 nm and the calibration curve of caffeine

To determine the amount of caffeine in solubilization study by using UV

spectrometry, first of all the calibration curve of caffeine was made by serial

0

0.05

0.1

0.15

0.2

0.25

0.3

A B C

conc

. of

chi

tosa

n (m

g/m

L)

A = Vivaspin 300 kDa B = Vivaspin 100 kDa C = Centrisart 20 kDa


- 34 -

concentration of caffeine as shown in Fig.17. The linear curve was achieved with R2

= 0.9989. Based on this calibration curve, the amount of caffeine which was

solubilized by chitosan could be calculated.

First of all the saturation concentration was determined as reference point for

caffeine solubility at 25°C. In Fig.18 cs is shown as dashed line at a concentration of

26.2 mg/mL which is in agreement with literature data [141]. The concentrations of

chitosan in contact with the saturated caffeine solution were then varied from 0.075

mg/mL to 0.30 mg/mL. Subsequently, using several concentration of chitosan ranging

from low (0.075 mg/mL) to high (0.3 mg/mL), the solubility of caffeine was influenced

by the concentration of chitosan. This trend was shown in Fig.18 that indicating

roughly a linear concentration between chitosan concentration and additional

dissolved caffeine amount (R2 = 0.9413). Hence, chitosan clearly has the capability

to solubilize caffeine.

3.4 Conclusions

Fourier transform infrared spectroscopy (FTIR), differential scanning

calorimetry (DSC) measurements and a solubilization study could demonstrate an

interaction between caffeine and chitosan. The elucidation of the interaction

betweencaffeine and chitosan which was already investigated by FTIR measurement

indicated that it was a complex formation and there was no peak shift of the carbonyl

R2 = 0.9413

25.0

26.0

27.0

28.0

29.0

30.0

31.0

0 0.05 0.1 0.15 0.2 0.25 0.3

conc of chitosan (m g/m L)

conc of caffe ine (m g/m L)

Cs

Fig. 18: Solubilization of saturated caffeine by adding chitosan. Dashed line represents the saturation concentration of caffeine (cs)


- 35 -

group at 1646 cm-1 and the methyl amine group of chitosan at 1020 cm-1. The

formation of such a complex was supported by DSC measurements where the glass

transition temperature revealed that molten caffeine is soluble in molten chitosan at

temperatures above 200°C. Furthermore, the changes observed in turbidity of the

saturated solution of caffeine becoming clear when adding chitosan was another

clear hint for this interaction taking place between caffeine and chitosan. UV

measurements further allowed quantifying the effect of chitosan on the solubility of

caffeine. Initially, without chitosan, the solubility of caffeine was 26.2 mg/mL in water

at room temperature. By adding chitosan up to 0.3 mg/mL the concentration of

caffeine solubilized was found to be 30.07 mg/mL. So, this amount of chitosan (0.3

mg/mL) could interact with caffeine and increase the saturation concentration by ~4

mg/mL.

IV. Nanoparticulate Formulation Intended for Hair F ollicle Targeting

- 36 -


4. 1. Introduction

Investigation of the use of drug carrier systems to treat hair loss is still in early

stage. In Human life, hair loss is a condition which occurs more frequent along with

time and increasing activities of human leading to stress conditions during daily work.

When a person experiences stress without realizing, so the stress condition lead to a

weakened metabolic system. Another reason for hair loss is because of the natural

aging process enhanced by hair care. After a hair is lost, usually the new hair can

grow again; if not baldness will occur [142,143]. Shampoo containing caffeine, as a

non-particulate formulation, has been available in commercial trade to stimulate hair

growth for quite some time [31,128,144]. In general, hair follicles were believed to be

a minor pathway for transdermal drug delivery into human skin [62,83,87,145] due to

the small hair follicle density of an average of 0.1% [38]. Based on recent

publications, it is known that particulate formulations show better penetration into hair

follicles than formulations without particle [62,63]. Particulate formulations, micro and

nanoparticles (NPs), which have been evaluated in several experiments about

transfollicular penetration indicated that particles with an optimal size of 320 nm can

penetrate deeply into the hair follicles [63]. However, for targeting of hair follicles,

particles size from 500 to 600 nm are believed to be well suited [86]. Beside the

possibility to penetrate into the hair follicles, the uptake into cells, especially

Langerhans cells, was according to literature observed for particles of around 40 nm

[146]. So depending on the specific need and target within the follicle different-sized

particles could be utilized.

To achieve a good drug carrier system the applied materials (polymers) have a

large impact. The utilization of natural and synthetic polymers is foremost to increase

the capability of carrier systems for drug delivery. Until now, some natural polymers

which have been successfully purified and synthetic polymers have been produced to

serve the formulation of NPs [107,147,148]. There are several requirements for these

polymers regarding regulative issues such as the biodegradability, biocompatibility

and benign character (toxicity) as well as technological aspects such as the

interaction between polymer and active pharmaceutical ingredient (API) influencing

the release profile from the NPs. Based on these criteria, the polymers which were

most extensively applied for creating nanoparticulate delivery systems over one


- 37 -

decade are poly(lactic-co-glycolic acid) (PLGA) and chitosan. NPs based on PLGA

were usually prepared by using emulsion solvent diffusion techniques which can be

considered the standard methodology. Nevertheless, PLGA NPs could also be

prepared by nanoprecipitation, emulsion coalescence and spray drying depending on

the specific goals of the formulation process [132,148-151].

It is also known that by using PLGA, a good encapsulation has been achieved

for hydrophobic APIs. In contrast, the encapsulation of hydrophilic drugs is not an

easy task and influenced by several factors such as kind of the polymer, the organic

solvents and correctly chosen stabilizers [108,115,152-154]. Nevertheless, some

studies reported, that by optimizing these factors, hydrophilic APIs could be loaded

into PLGA NPs by nanoprecipitation [102,103]. A combination of PLGA and chitosan

was chosen to vary the particle properties [110,111]. For delivery purposes, the

presence of amine functional groups of chitosan allowed to make use of the

electrostatic effect, of hydrogen bonds, and Van der Waals forces to interact with the

carbonyl functional group of the bioactive compound [155,156].

In this study, preparation of chitosan-PLGA NPs was performed by using

emulsion solvent diffusion method. Two types of homogenizer -ultrasonifier and

ultraturrax- were evaluated as well as two types of chitosan: Protasan UPCL 113 and

Kitozyme. In addition, PLGA NPs without chitosan were also prepared as a control.

Based on the literature reports that the double emulsion method for preparation of

NPs could load hydrophilic drugs, this method was also used for comparison. Pure

chitosan NPs without PLGA were prepared by using the crosslinker tripolyphosphate

(TPP). NPs were then characterized regarding size, polydispersity index, zeta

potential and morphology. The appropriate formula in terms of size distribution and

morphology was selected to be loaded with caffeine. Subsequently, NPs were

purified and % encapsulation efficiency as indirect EE of caffeine from the

supernatant was calculated. In terms of particles, the direct EE and loading of

caffeine was determined after NPs were dissolved. And finally, the release profile of

caffeine from the NPs was monitored.


- 38 -

4. 2. Materials and Methods

4.2.1. Materials

PLGA 50:50 (Resomer RG 503) with a molecular weight Mw of 24,000 - 38,000

Da was obtained from Evonik Industries (Darmstadt, Germany). Chitosan chloride

with a degree of 75% deacetylation (Protasan UPCL 113) with a molecular weight <

150,000 Da was obtained from Novamatrix (Norway), chitosan base (Kiomedine®)

with a degree of 75% deacetylation was obtained from Kitozyme (Herstal, Belgium).

Polyvinyl alcohol (PVA) (Mowiol 4-28) was obtained from Kuraray specialties

(Frankfurt, Germany), Pluronic F68 was obtained from BASF (Ludwigshafen,

Germany), Tripolyphosphate (TPP) was bought from Merck, Darmstadt, Germany,

cellulose membranes with a MWCO 12,000 – 14,000 Da were bought from Medicell

International LTD (London, UK) and caffeine anhydrous came from Sigma Aldrich (St

Louis, USA). For scanning force microscopy muskovite mica was obtained from

Plano Planet GmbH, Wetzlar, Germany. Silica wafers were obtained from Wacker

Chemie, Germany. NSC 16/50 non-contact silicon cantilevers, MikroMasch,

Cambridge, UK. All other solvents and chemicals were commercially available and of

highest grade.

4.2.2 Equipment

Zetasizer Nano ZS from Malvern Instruments, Worcestershire, UK was used to

determine the size and the zeta potential of the particles. The morphology of NPs

was determined by Atomic Force Microscopy (AFM) using a Nanoscope IV Controller

from Digital Instruments, Bruker Corporation, Billerica, USA and scanning electron

microscopy (SEM) using a EVO HD from Carl Zeiss, Jena, Germany.

4.2.3. Preparation of nanoparticles

The common method to form NPs loading hydrophilic drug is based on a

double emulsion[100] . It was formed by using 2.5 mL PVA 2.5 mg/mL containing

6.25 mg/mL caffeine which was dropped into 40 mg/mL PLGA in 2.5 mL ethyl

acetate. The mixture was then placed in an ice bath under stirring for 1 hour at 750

RPM. To produce a nanoemulsion, ultraturrax at 13,200 RPM was applied for 10

minutes. The second emulsification step was formed by using Pluronic F68 as


- 39 -

stabilizer with concentrations of 15 and 20 mg/mL (2.5 mL for each). The resulting

emulsion was stirred over night at 500 RPM to evaporate the organic solvent.

Emulsion solvent diffusion method was used to prepare the chitosan-PLGA

NPs as described before [110,111]. In brief, the emulsion was prepared by using

PLGA 40 mg/mL in 2.5 mL ethyl acetate as organic phase. The aqueous phase

consisted of 2.5 mL of 25 mg/mL PVA solution, chitosan 0.3 mg/mL, and 6.25 mg/mL

caffeine. The aqueous phase was dropped into ethyl acetate as organic phase. The

mixture was placed in an ice bath under stirring for 1 hour at 750 RPM. To produce a

nanoemulsion, both ultrasonifier at 10% amplitude (A) with 500 Joule for 1 minute

and ultraturrax at 13,200 RPM for 10 minutes in an ice bath were applied. The

resulting emulsion was stirred over night at 500 RPM to evaporate the organic

solvent. For size comparison, blank PLGA NPs and chitosan-PLGA NPs were

prepared without caffeine using the same protocol as for the single-step emulsion

solvent diffusion. Besides Protasan, another type of chitosan - Kiomedine originating

from mushrooms and produced by Kitozyme was also used for comparison. Pure

chitosan NPs without PLGA core were also prepared by using ionic gelation method

based on the crosslinker tripolyphosphate (TPP). Chitosan (Protasan) was dissolved

in demineralized water to make a stock solution of chitosan (0.625 mg/mL) and TPP

was dissolved also in demineralized as stock solution of TPP (0.125 mg/mL).

Chitosan NPs were formed by adding 2 mL TPP 0.125 mg/mL into 2 mL chitosan

0.625 mg/mL. The resulting mixture was stirred for 4 hours.

4.2.4. Purification of nanoparticles and %EE of caffeine

5 mL suspension of NPs was placed in Vivaspin-20 (Sartorius, Göttingen,

Germany). The suspensions were purified by using a centrifuge at 14,000 x g at

T = 8°C for 30 minutes. The supernatant were collected to determine the

encapsulation efficiency (%EE) of caffeine by using HPLC-analysis. The mobile

phase of HPLC was a phosphate buffer pH 2.6 and acetonitrile (90:10) with a flow

rate of 1.2 mL/min. The HPLC column was a reverse phase column C18 with

controlled temperature at 25°C.

The amount of NPs was obtained from lyophilized samples. After that, this

sample allows to quantify the loading of drug in the formulation. The loading can be

calculated based on the amount of drug in NPs and the weight of the samples used


- 40 -

for investigation. For lyophilization, sample was pre-frozen at T = -80°C and then the

drying process was continued by a conventional Alpha 2-4, freeze-dryer (Christ,

Osterode, Germany).

4.2.5. Size, PDI and zeta potential and morphology study of the nanoparticles

Determination of size, PDI and zeta (ζ) potential were the common

approaches to determine the properties of the NPs. After evaporation of the organic

solvent, the NP suspensions were characterized regarding size, ζ-potential, and

polydispersity index (PDI) using dynamic light scattering (DLS) from Malvern

Instruments, Worcestershire, UK. NPs were purified by Vivaspin20 and the filtered

NPs were washed with 0.2% trehalose which also acted for the pellet redispersant.

The redispersed NPs were analyzed regarding their morphology. The morphology of

NPs was visualized by AFM and SEM measurements. Before the measurements,

NPs were dilluted in demineralized water using vortex and dried overnight at room

temperature. AFM measurements were done with a Bioscope with a Nanoscope IV

controller (Veeco Instruments, Bruker, Germany). Samples were prepared by

dropping 20 µL of diluted sample on freshly cleaved mica. The NPs were investigated

after drying under ambient condition with tapping mode using a tip with cantilever of k

= 40 N/m and a resonance frequency of ~250 kHz. For SEM measurements, the

samples were dried under ambient conditions and coated with gold using a Quorum

Q 150 ES sputter coater at 40 mA for 50 s in order to render the sample conductive,

improving the image quality (thickness of the gold coating was less than 20 nm). For

visualization a ZEISS EVO HD 15 was used in high vacuum mode, applying an

acceleration voltage of 5 kV.

4.2.6. Dissolving nanoparticles and determination of loading

Several organic solvents such as ethyl acetate, dichloromethane,

dimethylformamide (DMF), tetrahydrofurane (THF) and dimethylsulfoxide (DMSO)

were tested to dissolve chitosan-PLGA NP. As a control, PLGA NPs were used in

this experiment. PLGA dissolves in DMSO and ethyl acetate and chitosan is only

soluble in acidic solution. DMSO is soluble in water but ethyl acetate is only partially

soluble in water. The NPs were placed into 0.1% acetic acid, followed by DMSO to

obtain a one-phase system. This approach required heat treatment at T = 60°C for 9


- 41 -

hours and 15 minutes sonication successively. Afterwards, the amount of caffeine

loaded was determined (direct method).

4.2.7. In vitro release study

For the release study, Franz Diffusion Cells were set up. For the study the

same volume of acceptor and donor compartment was used. The drug can diffuse

effectively into the acceptor compartment and the speed of stirrer bar also the sink

condition of each cell were important factors for release study using Franz Diffusion

Cell. Performing release study followed the procedure described before [157] . The

donor compartment was filled with 40 mg lyophilized NPs containing caffeine

suspended in 0.5 mL demineralized water. A Medicell membrane with MWCO 14,000

Da was placed in between donor and acceptor compartment. The acceptor

compartment was filled with 12.5 mL of phosphate buffered saline (PBS) pH 7.4. The

caffeine released to the acceptor compartment at room temperature was sampled

using a long syringe; a Pasteur pipette was used to refill the acceptor compartment

through the sampling port. The amount of released caffeine was determined by

HPLC analysis. In this determination, mobile phase phosphate buffer pH 2.6;

acetonitrile = 90:10 was used, the retention time was 5.1 min, at λmax was 262 nm.

The shift of λmax from 272 to 262 nm due to the influence of acetonitrile and

phosphate buffer in mobile phase of HPLC. Flow rate was 1.2 mL/min and injection

volume 20 µL. For comparison, the passive diffusion of pure caffeine and from a

mixture of caffeine-chitosan was also determined.

4.3 Results and Discussion

4.3.1 Size, PDI, and zeta potential of the prepared NPs

After we established the binding capacity of chitosan to caffeine, we

developed a formulation with NPs loading caffeine using biodegradable and

biocompatible polymers (chitosan and PLGA) by using the established solvent

diffusion evaporation method (single emulsion).

The common type of drug to load PLGA NPs is hydrophobic. Nevertheless,

hydrophilic drugs can also be loaded into PLGA NPs using double emulsion[102,


- 42 -

103] . Therefore double emulsion was also used to encapsulate caffeine as reference

method.

Both methods -single and double emulsion- have used PLGA as polymer due

to its well-known properties. Resomer RG 503 is a PLGA which has 50% glycolic and

50% of lactic acid groups and a molecular weight Mw of 24,000 to 38,000 Da. Based

on literature, PLGA in combination with chitosan is able to form PLGA NPs (core)

coated with chitosan. This system has been described to be a good drug carrier

system [110, 111, 115, 158]. In addition, chitosan-PLGA NPs prepared using the

standard method (single emulsion) were described showing better PDI than other

approaches. Besides the formulation of hybrid particles based on PLGA and

chitosan, chitosan NP could also be formed by combining it with alginate, gelatine

and crosslinker molecules. Furthermore, pure chitosan particles were prepared using

the crosslinker tripolyphosphate (TPP) which was chosen due to its good safety

profile.

0

50

100

150

200

250

300

350

400

A B C D ESamples

Size (nm)

-25

-15

-5

5

15

25

35

45

Zeta Potentia l (mV)

Figure 19. Size and charge of PLGA, chitosan-PLGA, and TPP-chitosan particles. The dashed

line represents a guide to the eye to show the reference size from pure PLGA NP.

First of all, double emulsion as common method to load polymeric particles

with hydrophilic drugs was used to prepare PLGA NPs loading caffeine. This

preparation method was then compared to a single emulsion preparation containing

PDI 0.21

PDI 0.06

PDI 0.16

PDI 0.17

PDI 0.02

A. PLGA NP B. Double Emulsion with 1.5%Pluronic and 2% PVA C. Double Emulsion with 2% Pluronic and 2% PVA D. Chitosan-coated PLGA NP E. Chitosan-TPP NP


- 43 -

forming chitosan-PLGA NPs using the specific interaction between chitosan and

caffeine for loading. As shown In Fig.19 size and ζ-potential of NPs prepared by the

double emulsion methods with two concentrations of the stabilizer Pluronic F68 are

depicted. The two double emulsion formulations resulted in particles smaller than

300 nm and with a PDI of 0.16 indicating a relatively broad size distribution of the

carriers. However, single emulsion method for chitosan-coated PLGA NPs showed

smaller sizes (d ~ 250 nm) and a very good PDI of ~0.06. As control, blank PLGA NP

were prepared yielding the smallest and most well distributed particles with sizes of

208 nm and a PDI of 0.02. Particles prepared by the ionic gelation method (chit-

TPP), using tripolyphosphate (TPP) showed sizes of more than 300 nm and the worst

size distribution (PDI = 0.21) (Fig.19). Looking a bit closer at the size distribution

(Fig.20) the chit-TPP NPs had the largest range with a full width at half maximum

(FWHM) of 205.49 nm. The second largest FWHM was found for PLGA NPs

prepared by double emulsion with particles showing a FWHM of 166.38 nm, followed

by chit-PLGA NPs with FWHM = 155.25 nm and the control PLGA NPs with the

smallest distribution (FWHM = 123.75 nm).

-5

0

5

10

15

20

25

30

35

0 200 400 600 800

Size (nm )

Intens ity (%)

chit-TPP NPs

chit-PLGA NPs

PLGA NPs

Double emulsion PLGA NPs

Figure 20. Size and distribution of chit-PLGA, PLGA, chit-TPP and PLGA double emulsion

The zeta potential of the different preparations followed the expected trends

displaying negative ζ-potentials for the preparations without chitosan and positive ζ-

potentials for the chitosan containing formulation because of the amine groups of

chitosan (Fig.19).

Broad distribution

FWHM 155.25 nm


- 44 -

Based on the results of the characterization of NPs size, PDI and ζ-potential,

especially chitosan-TPP NPs did not show good properties with a too broad

distribution (PDI = 0.21). Therefore, these particles were not further investigated. The

preparation of PLGA NP and the chitosan NP was further evaluated with respect to

the dispersion method and the type of chitosan used.

Table 1. Ultraturrax and Ultrasonifier as Homogenizer

Ultraturrax Ultrasonifier Samples Size (nm) PDI ζ−Pot (mV) Size (nm) PDI ζ−Pot (mV)

PLGA NP

208.2 ± 1.91 0.042 ± 0.002 -10.31± 0.045 202.6 ± 1. 104 0.036 ± 0.001 -10.01 ± 0.038

Chit-PLGA NP

267.8 ± 4.49 0.081 ± 0.014 17.72 ± 0.048 253.1 ± 2. 621 0.066 ± 0.012 17.94 ± 0.042

Standard deviation of (n=3)

The effect of homogenizing method on the properties of NPs is shown in

Tab.1. There was no difference with respect to size and PDI found. However, using

the ultraturrax the 10 minutes preparation time caused an increase in temperature to

20°C with ice bath and even to 42°C without ice bath. On the other hand, ultrasonifier

used for 1 minute only showed an increase in temperature to 11°C with ice bath

(without ice bath was 18°C) resulting also in NPs with very good particulate

characteristics. Since this method was less time consuming while providing a

reduced temperature effect and both methods gave comparable size distributions the

method based on the ultrasonifier was chosen for further experiments.

Table 2. Impact of the kind of chitosan (Protasan and Kitozyme) on the NP properties

Characterization Samples Size (nm) PDI ζ−Pot (mV)

Protasan 255.5 ± 2.718 0.068 ± 0.015 17.40 ± 0.062

Kitozyme 276.5 ± 4.301 0.092 ± 0.060 14.32 ± 0.091 Standard deviation of (n=3)

As there is more than one kind of chitosan - Protasan and Kitozyme- the

impact of the two types of chitosan was studied. Protasan was able to produce NPs

with better PDI than those prepared with Kitozyme (Tab.2). Overall the differences

were small. However, one important disadvantage of Kitozyme is its solubility profile

restricting the preparation. The preparation of Kitozyme solution was more time

consuming because to dissolve this chitosan in acidic pH needs sonication for 2


- 45 -

minutes followed by stirring for 15 minutes. In contrast, Protasan dissolved

immediately in demineralized water. To keep the temperature change of the process

low, an ice bath was needed. In addition, the solubility of caffeine is also reduced by

reducing temperature.

Hence lower preparation temperatures seemed to be meaningful for

preparation as caffeine solubility changes. Taken all these data into consideration,

chitosan-PLGA NPs which were formed by single emulsion method using Protasan

and an ultrasonifier to form the nanoemulsion were chosen for drug loading. All

further investigations as determination of the encapsulation efficiency, the loading

capacity and the release of caffeine were performed with this formulation.

4.3.2. Purification of nanoparticles, %EE of caffeine and morphology study

Supernatant after purification of chitosan-PLGA NPs loading caffeine was

collected. The amount of caffeine in the supernatant was quantified by using HPLC-

analysis. EE was calculated based on the following formula :

EE of caffeine from chitosan-PLGA NPs was found to be (19 ± 1.1)%. In

comparison the EE of caffeine in PLGA NPs prepared by the double emulsion

method was only (14 ± 1.1)%. Obviously, the addition of chitosan forming chitosan

NPs using single emulsion led to an increased EE compared to the double emulsion

approach.

Based on the AFM and SEM images it could be demonstrated that the

morphology of chitosan-PLGA and PLGA NPs was spherical with a smooth surface

(Fig. 21). From Fig.21, analysis of the average size of NPs was conducted by using

Section Analysis of the Nanoscope software. Average size of NPs from AFM image

(Fig.21a) of chit-PLGA NPs was 215.31 nm and the PLGA NPs (Fig.21b) were 147.8

nm.

The fact that the chit-PLGA NPs were larger in size was expected due to

previous results [110] and the increased viscosity of the aqueous phase due to the

EE = total amount of caff - amount of caff in supernatant x 100% total amount of caff

caff is caffeine


- 46 -

presence of chitosan. Analysis of the average sizes of NPs using SEM images, chit-

PLGA NPs (Fig.21c) yielded an average of 164.7 nm and the blank PLGA NPs

(Fig.21d) were found to be 135.5 nm by using ImageJ. All the NPs from SEM images

were smaller than the hydrodynamic sizes determined by light scattering because of

the drying and vacuum process. This also explains the differences between SEM

measurements, performed in vacuum, and AFM measurements done under ambient

conditions. In addition, for AFM measurements the unknown effect of the tip

geometry is always present increasing slightly the object size. Nevertheless, the

particle sizes are all in the same size range within the variations due to the

techniques and support the obtained narrow size distribution.

Chitosan-PLGA NPs PLGA NPs

A

B

C

D

Figure 21. AFM images of chit-PLGA NPs (A), PLGA NPs (B) and SEM images of chit PLGA (C) and PLGA NPs (D).


- 47 -

4.3.3. Dissolving of chitosan-PLGA nanoparticles to determine the loading

To determine the amount of drug incorporated in the particles it is necessary

to dissolve the particles, evenly disperse the drug and then separate the drug from

the other components for analysis. Therefore, the dissolution step of the particles is

essential to evaluate directly the amount of loaded drug.

After purification and lyophilization of chitosan-PLGA NPs the samples were

obtained as powders. To suspend them, demineralized water was added followed by

vortexing. By this step, we obtained the NPs well suspended as a stable suspension.

These suspensions were used for the dissolving step. However, dissolving the NPs is

not always an easy task. For instance, in literature is already described that even

plain PLGA NPs do not always completely dissolve by using good organic solvents

such as DMSO [159]. Having this in mind, it was not surprising that the hybrid PLGA

NPs with chitosan and stabilizers [160] were more difficult to dissolve. Chitosan,

representing the outer layer of the particles [111], is well soluble in acidic solution.

0

0.5

1

1.5

2

2.5

3

3.5

263 363 463 563 663 763

wavelenght (nm)

A

dispersed particles

dissolved particles in one phase

Figure 22. Turbidity measurement of chit-PLGA NP suspension (grey line) and the absorption signal (≙ light scattering) after dissolving chit-PLGA NPs (black line).

Therefore an approach based on an immersion in an acidic solution was used

first. An acetic acid solution 0.1% at T = 60°C for 9 hours was applied, followed by 15

minutes sonication. Afterwards, DMSO was added to the system to obtain an one-

phase system of drug and polymeric entities (particles were completely dissolved).

The successful dissolution of the NPs was investigated by turbidity measurement

using UV/Vis spectroscopy. Before dissolution, the particles dispersed in water

clearly showed a scattering signal (upper curve in Fig.22). After acetic acid-DMSO


- 48 -

treatment, the signal strongly drops indicating completely dissolved chitosan-PLGA

NPs because of reduced or no colloidal scatteres. Light scattering data of the

dissolving NP showed clear solution (no measured absorption signal in the range

from 360-800 nm). This result showed that the problem regarding dissolving

chitosan-PLGA NPs could be solved using such a two-step dissolution process. After

knowing the condition to dissolve chitosan-PLGA NPs, we could continue to

determine the amount of successfully loaded caffeine into the NPs. The solution of

dissolved NPs was centrifuged at 14,000 x g for 90 minutes to remove large objects

and filtered with Chromafil filter 0.2 µm to avoid agglomerated material eventually

present. Afterward, 100 µL of this solution was diluted with 900 µL mobile phase

(buffer phosphate pH 2.6; acetonitrile = 90:10). Detection wavelength was 262 nm

with retention time of 5.1 min and flow rate was 1.2 mL/min with injection volume of

20 µL. The amount of caffeine was determined and the %EE by this direct method

was found to be (12.33 ± 1.7094)% based on the following formula :

The loading of caffeine was 0.059, indicating that in 100 mg NPs 5.9 mg caffeine

were contained. The loading was calculated based on the following formula

Based on this result, the %EE and the loading is low as typical of hydrophilic drug.

However this values show that chit-PLGA NPs can load caffeine. Due to more

specific localization of NPs in hair follicles, the small amount of NPs is sufficient.

4.3.4. In vitro release study

Franz Diffusion Cells were prepared to perform release studies according to a

method which was described previously [161]. For this approach, lyophilized NPs

loading caffeine were suspended in demineralized water and the caffeine released to

the acceptor compartment at room temperature was monitored [157]. The sampling

Loading = amount of caffeine in NPs total weight of NPs

%EEdirect = amount of caffeine in NPs x 100% amount of caffeine added in the beginning


- 49 -

was performed by a long syringe and Pasteur pipettes were used to refill the acceptor

compartment through the sampling port. The membrane with MWCO 14,000 Da was

placed in between acceptor and donor compartment. The concentration and hence

the amount of released caffeine was determined by using HPLC analysis.

The influence of chitosan-PLGA NP loaded caffeine led to a prolonged and

slower release of caffeine from the particles as shown in Fig.23. Up to 24 hours, 40%

caffeine was rapidly released (burst release) and the remaining amount of caffeine

was continuously released until at 56 h a plateau was reached. The missing amount

of caffeine should be still associated with chitosan. Based on literature the

encapsulation of drug into NPs can influence the pattern of drug release [162].

Obviously, chitosan coating PLGA NPs could reduce the burst release of caffeine

from the particles. This is because of the interactions between caffeine and chitosan

as shown in chapter 3. Furthermore, according to recent reports chitosan was found

after freeze drying changing its swelling behavior. Chitosan was observed to show a

time dependent hydration and swelling. As consequence the release of caffeine

incorporated in such a swollen chitosan layer would be prolonged compared to the

non-entrapped caffeine. Pure caffeine, as expected, showed the fastest passive

diffusion across the membrane up to 6 h and achieved 75% released amount. For

the mixture of caffeine-chitosan, passive diffusion of caffeine was slower in the first 6

hours than the diffusion across the membrane for pure caffeine but faster than the

caffeine diffusion from the chit-PLGA NPs. After 24 hours the maximum of permeated

caffeine was achieved at ~ 70 %. This was the same level as found for free caffeine

and represents the equilibrium level. The slower permeation across the membrane

from the chit-PLGA NP might be due to the particulate nature. In addition to the

interaction between chitosan and caffeine the geometric restrictions of the chitosan

layer at the particle interface together witht the swelling will contribute and further

slow down the release from the particles. This permeation profile across the

membrane indicated that caffeine was interacting with chitosan and that the resulting

complex (caffeine-chitosan or caffeine-chit-PLGA NP) slowed down release and

hence the free diffusion of the drug.


- 50 -

0

10

20

30

40

50

60

70

80

90

0 6 12 18 24 30 36 42 48 54 60Time (h)

% release

chit-PLGA NP

caffeine

caffeine-chitosan

Figure 23. Release/diffusion profiles of caffeine across a membrane for chit-PLGA NP loading caffeine, pure caffeine and the physical mixture of caffeine and chitosan.

4.4 Conclusions

In this study, we propose a new biodegradable nanocarrier for loading the

hydrophilic drug caffeine. The NPs which were prepared showed spherical shape

with sizes of 208±1.104 nm for pure PLGA, 254±2.621 nm for chitosan-PLGA, and

351±12.051 nm for TPP-chitosan. The polydispersity index of the two NP

preparations based on PLGA was better than that of the ionic gelated chitosan.

Therefore, this preparation method was not further investigated. The zeta potential of

the NP based on PLGA was found to be negative whereas the chitosan-containing

NPs were, as expected, positive. To load the hydrophilic drug the double emulsion

approach and the single emulsion set-up were used and compared. The

encapsulation efficiency of caffeine to the chitosan-PLGA NPs was determined to be

(19 ± 1.1)%. It was higher than that found for the double emulsion method without

chitosan (only ~14%). Thus the interaction between caffeine and chitosan enabled a

higher loading. Therefore, this system seems to be better suited for an applicaton

targeting hair follicles. The loading of caffeine into chitosan-PLGA NP could also

prolong the release of caffeine compare to the passive diffusion of caffeine and

mixture of caffeine-chitosan. This reduced release of the drug could further support a

reservoir effect of the particles when accumulating in hair follicles.

V. Photoacoustic Microscopy to Image Model Hair Fol licle

- 51 -


5.1 Introduction

To bring drugs across the skin is one of the biggest challenges for a drug

delivery system [163-165]. The optimization of such systems is difficult, because skin

is well known as a good barrier regarding to the dense, poorly permeable

corneocytes and overlapping alignment of the cornified cell layers of skin. Another

difficulty is that not all drugs could overcome the epidermis by penetration. It is

dependent on the partition and diffusion coefficient of the active pharmaceutical

ingredient (API). APIs with high partition coefficient can pass through the stratum

corneum (SC) and for APIs with low partition coefficient it is difficult to penetrate SC.

Molecular weight (MW) of the API plays also an important role for skin penetration.

500 Dalton (Da) is the molar mass often stated as exclusion limit for dermal

penetration and permeation through the SC [38]. Against these difficulties, the

alternative strategy is to collect the API in the hair follicle as an intriguing approach

gaining more attention for targeting API because the hair follicles are known as an

efficient storage and penetration pathway for topically applied substances [145].

Follicular penetration for targeting with NPs is a relatively new topic of research.

Using this pathway, the NP loaded with API can enter the hair follicles and then

release the API without need to disturb the skin barrier (non-invasive delivery)

[87,92,145].

Visualization of sectioned skin could be performed by using confocal laser

scanning microscopy (CLSM). However imaging entire hair shafts in skin using

CLSM faces drawbacks due to the size and the tilted orientation of the hair shaft in

the skin [166]. Nevertheless, an imaging of intact hair follicles would be necessary

and helpful to identifying the presence and location of the NPs. Usually sectioned

skin was used for CLSM but finding intact hair follicles is a time consuming task by

this technique. On the other hand, photoacoustic microscopy allows visualizing such

large scale structure non-invasively [167-169]. A necessity for sufficient quality of

images of photoacoustic microscopy is the usage of a contrast agent such as

magnetite (iron oxide) [170]. Iron oxide can absorb the near infra red light and emit

ultrasound. This ultrasound waves can then be detected and converted to an image

by using an ultrasound-transducer [171,172]. Recently, researchers have focused to


- 52 -

load iron oxide into NP made from biodegradable polymer such as dextran, gelatin

and PLGA to obtain paramagnetic properties of the NPs. These properties are

especially advantageous with respect to cleaning and collecting of NPs

[169,173,174]. The polymer of choice in this study is the synthetic polymer poly

(lactic-co-glycolic acid) (PLGA) which is a well-known biodegradable and

biocompatible pharma polymer [96,175,176]. This property is a basic requirement for

therapeutic usage of NPs [44,99,106]. Thus magnetite is intended to be incorporated

into PLGA NPs drug carrier.

Another advantage for the application of NP as drug delivery system is the

reduction of undesired side effects of the API [177-179]. A controlled size distribution

is a central requirement for a pharmaceutical product [180-182]. Stronger restrictions

are laid upon the selection of the methods that will be used for NP production,

avoiding harmful substances during the production process. Additionally, the stability

of NP should be considered for the delivery system [183,184].

For the preparation of NPs, the emulsion solvent diffusion and

nanoprecipitation are known as common methods [100,102,184]. Emulsion solvent

diffusion and nanoprecipitation methods were used to prepare NPs. The effect of

magnetite concentration on the formulation was investigated, in terms of the physical

properties of NPs. Hydrodynamic size, dispersity index and zeta potential were

determined by dynamic light scattering (DLS). The morphology of NPs was

determined by scanning electron microscopy (SEM) and atomic force microscopy

(AFM). The localization of magnetite in the PLGA NPs was determined by AFM and

transmission electron microscopy (TEM). Furthermore, the magnetite-PLGA NPs

were applied for imaging of the model hair shaft by using photoacoustic microscopy.

5.2. Materials and Methods

5.2.1. Materials PLGA Resomer RG 503 (50:50) was obtained from Evonik Industries. Its

molecular weight is 24,000 - 38,000 Da. Polyvinyl alcohol (PVA) Mowiol was obtained

from Kuraray, Pluronic F68 was obtained from BASF, agarose and oleic acid-

stabilized magnetite in toluene were purchased from Sigma Aldrich,

aminohexanamine-stabilized magnetite in dimethyl formamide (DMF) was obtained

from the Department of Physical Chemistry, Saarland University. For scanning force

microscopy, as sample support muskovite mica was obtained from Plano Planet


- 53 -

GmbH, Wetzlar, Germany. Silica wafer was obtained from Wacker Chemie,

Germany. NSC 16/50 non-contact silicon cantilevers from MikroMasch, Cambridge.

Syringes (Sterican-26G) with diameter of 0.45 mm were obtained from Braun Medical

AG, Emmenbrücke, Germany. All other solvents and chemicals were from the

highest grade and commercially available.

5.2.2. Equipment

Zetasizer Nano ZS from Malvern Instruments, Worcestershire, UK. Atomic

Force Microscope (AFM) with a Nanoscope IV (Digital Instruments), Bruker

Corporation, Billerica, USA. Scanning electron microscopy (SEM) using a SEM EVO

HD 15 from Carl Zeiss and sputter Quorum Q 150 RES from Judges scientific, UK.

Transmission Electron Microscopy (TEM) JEM-2010, JEOL GmbH, Eching,

Germany. Confocal laser scanning microscope ZEISS LSM 510 META, Carl Zeiss,

Jena, Germany. Optoacoustic signals of pulsed Nd-YAG laser system (H7000) was

provided an optical microscope (Olympus IX 81), Tokyo, Japan.

5.2.3 Preparation of PLGA NPs loaded with aminohexanamine-stabilized magnetite

To prepare nanoparticles containing magnetite, nanoprecipitation method

[185] was used. 25 mg/mL PLGA and 1 mg/mL of magnetite particles were dispersed

in acetone under stirring for 30 minutes. Nanoprecipitated particles occurred when

this organic phase was immersed to an aqueous phase contained Pluronic F68 as

stabilizer. The immersion process was conducted by using a syringe. Then acetone

was evaporated overnight at room temperature (~22°C).

To compare with nanoprecipitation another method for the preparation of NPs

-the single emulsion solvent diffusion- was udsed [116,169]. In brief, 25 mg/mL PLGA

was dissolved in ethyl acetate as organic phase. After that, 1 mg/mL

aminohexanamine-stabilized magnetite in DMF was added to 6 mL of the organic

phase under stirring followed by sonication for 30 minutes and vortex 2 minutes to

achieve a homogenous suspension. To form an emulsion, the organic phase was

dropped to the aqueous phase containing 2.5% PVA as stabilizing agent and stirred

for 1 hour. By using an ultrasonifier with 500 J for 30 seconds, the emulsion was

homogenized to form sizes in nm scale. Afterwards, dilution by adding 6 mL

demineralized water allowed ethyl acetate to mix with the aqueous phase. After that,

ethyl acetate was evaporated overnight at room temperature (~22°C). The


- 54 -

preparation was conducted by using several concentrations of magnetite: 0.5, 1, 2, 4

and 6 mg/mL.

To homogenize aminohexanamine-stabilizing magnetite into organic phase in

presence of DMF, several steps were needed. Regarding this limitation, in a second

approach, it was tried to remove and exchange DMF. Thus, aminohexanamine

stabilizing-magnetite in DMF was diluted in isopropanol and then poured into ethyl

acetate containing 25 mg/mL PLGA. The supernatant was separated from

precipitated magnetite-PLGA particles. The precipitate was dispersed in 2.5 mL ethyl

acetate. Once dispersed, the single emulsion method was conducted. Organic phase

was added drop by drop by using peristaltic pump into the aqueous phase containing

2.5% PVA. To emulsify this mixture, ultrasonifier with 500 J for 30 seconds was

applied, followed by dilution with 6 mL demineralized water. Evaporation of the

excess organic solvent was conducted at room temperature (~22°C) over night.

5.2.4 Preparation of PLGA NP loaded with oleic acid stabilized-magnetite

In the previous step, the influence of removing DMF was removed. In this step,

for comparison, the preparation of NPs was performed with oleic acid-stabilized

magnetite in toluene which is miscible with ethyl acetate. NPs were prepared by

using 25 mg/mL PLGA in ethyl acetate with 0.5, 1 and 2 mg/mL of oleic acid-

stabilized magnetite. The organic phase was mixed by stirring and then dropped into

PVA 2.5% as aqueous phase by using peristaltic pump. Stirring was continued for 1

hour. To form nano-sized NPs, homogenization by using ultrasonifier with 500 J for

30 seconds was applied. After that, dilution was conducted by adding 6 mL

demineralized water. Over night evaporation was applied at room temperature

(~22°C) to remove ethyl acetate.

5.2.5 Purification and characterization of nanoparticles

For purification, the suspension of NPs as much as 5 ml was placed in

Vivaspin-20 with a MWCO of 300,000 Da from Sartorius. The suspensions were

purified by using a centrifuge at 14,000 x g, T = 8°C, for 30 minutes. Filtered NPs

were washed and redispersed with demineralized water. For determination of the

physical properties of resuspended NPs, they were characterized regarding size,

polydispersity index (PDI) and zeta potential by dynamic light scattering (DLS) with a

Zetasizer Nano ZS (Malvern Instruments, Malvern, UK). Morphologic characterization


- 55 -

was performed by SEM an AFM. To achieve an adequate concentration of NPs for

the microscopic measurement, suspension was diluted 1:100 in demineralized water.

Then, for SEM measurement, 20 µL of the diluted sample was dropped on a silica

wafer and dried under ambient conditions. The sample was then coated with a layer

of ≤ 20 nm gold (sputter coater Quorum Q 150 ES at 40 mA for 50 s) in order to

render the sample conductive and improve image quality. For visualization a ZEISS

EVO HD 15 SEM was used in the high vacuum mode, applying an acceleration

voltage of 5 kV.

To obtain 3 dimensional information on the sample (20 µL of the diluted sample was

dried on the freshly cleaved mica) an atomic force microscope (Bioscope SPM) with

a Nanoscope IV controller (Digital Instruments, Bruker) was used. The NPs were

investigated under ambient conditions in tapping mode using a tip with cantilever of k

= 40 N/m, resonance frequency of ~250 kHz. After that, the AFM images were

analyzed with the AFM software (Nanoscope IV version 5.12R.3).

5.2.6 Determination of the magnetite core by using TEM, elemental analysis and AFM phase imaging

Determination of magnetite core incorporated into PLGA NPs was conducted

by TEM, elemental analysis and AFM phase imaging. TEM sample was prepared by

dropping the 20 µL of magnetite-PLGA NPs suspension on a copper grid. The

sample then was dried under ambient conditions on the surface of the grid. The

visualization was conducted by JOEL Transmission Electron Microscopy (model

JEM-2010 instrument, JEOL GmbH, Germany). The size of the magnetite NPs was

determined from the contrast (black spots) at the TEM image. Furthermore,

elemental analysis allowed the identification of the materials which were incorporated

into PLGA particles.

5.2.7 Preparation of model hair shaft

The idea was to create a void structure (mimicking the hair shaft) which was

then filled with PLGA NPs loading magnetite. Therefore a syringe was immersed into

1.75 % agarose. After the agarose solidified, a void was obtained by withdrawal of

the syringe. The model is composed of three layers (Fig.24). First was a based layer

(as a foundation), second was a gel forming layer containing the void, and the third

was a cover layer (to restrict diffusion of NPs during the CLSM and photoacoustic

measurement). First of all, to form the based layer, 5 mL agarose was poured into


- 56 -

Petri dish. After that, to form the gel layer, 12 mL agarose was added above the

based layer. In this layer a syringe containing the suspension of NP was immersed

before agarose could become solid. The model hair shaft was obtained when the

agarose became solid around the syringe. After that, the suspension of NP was

injected slowly into the void while the syringe was removed from the agarose.

Therefore the void was filled with the NPs suspension. The whole surface was then

covered with a thin layer of agarose. As control, pure magnetite and blank PLGA NPs

were used to perform photoacoustic imaging. Whereas for the CLSM imaging, the

PLGA NPs labeled with fluoresceinamine were used.

Figure 24. Sketch of the reparation of model hair shaft containing magnetite-PLGA NPs

5.2.8 Imaging model hair shaft by using CLSM and Photoacoustic Mcroscopy

For CLSM measurement using a confocal laser scanning microscope ZEISS

LSM 510 META, Carl Zeiss, Jena, Germany, agarose gel samples were sliced in

cubic blocks and placed on the cover slip. The slice was placed in the microscope

with the model hair shaft being parallel to the objective. This preparation was

necessary to omit the limitations regarding the focal distance of the objective. The

excitation was performed by using λ = 488 nm. The fluorescence signal was

collected after a band-pass 522/35.

Optoacoustic imaging is a selectively new method that relies on a good

absorbtion of light (in near infrared region). The sample is irradiated by short laser

pulses in the sub-nanosecond range generating broadband acoustic signals as a

result of the thermoelastic effect. These signals propagate through the sample and

can be acquired with adequate ultrasound transducers. When compared with

Zoom

Syringe containing magnetite-PLGA NP

Petri Disc containing 1.75% agarose

Gel layer

Model hair shaft containing magnetite-PLGA NP

Top layer

Gel layer

Top layer

Based layer


- 57 -

conventional ultrasound imaging, optoacoustic techniques have the advantage of

higher contrast since optoacoustic signal amplitudes are proportional to the local

absorption coefficient µa. Since the absorption is dependent on the concentration of

absorbing NPs, this effect can be used in order to assess the suitability of different

particle types (magnetite, magnetite-loaded polymer) as contrast agents.

For assessing the detectability of the different NPs types, an optoacoustic

microscope has been used. Photoacoustic microscopy setup was developed by

Fraunhofer Institute for Biomedical Technology (IBMT), ST Ingbert, Germany.

Optoacoustic signals were generated by a pulsed Neodymium-doped Yttrium

Aluminum Garnet (Nd-YAG) laser system (H7000). The data were amplified using a

single channel hardware platform (AMI-US/OA, Kibero GmbH) to obtain the

images[186] . For imaging of the prepared samples (model hair shaft loaded with

different NPs), two different optoacoustic imaging systems were used. The device

developed by Fraunhofer IBMT is based on an inverted optical microscope (Olympus

IX 81, Tokyo, Japan). It can be used as conventional optic microscope, acoustic

microscope or optoacoustic microscope. For photoacoustic microscopy, the

measurement could directly be conducted from the Petri dish containing the gel and

the model hair shafts.

For acquisition of optoacoustic signals, an acoustic detection unit has been

mounted to the microscope. It is based on a 2D piezo-scanner that allows point-wise

acquisition of signals from the investigated sample. Signals can be acquired using

focused high frequency ultrasound transducers. For achieving the highest resolution,

a 400 MHz ZnO transducer with a lateral resolution of 2.5 µm developed by

Fraunhofer IBMT is used [170]. The scanner and the transducer are attached to a

rotating column so that a fast exchange between the optoacoustic/ultrasound

modality and the conventional optical imaging mode is guaranteed. This is especially

helpful when optical microscopy was used to assist the position of targeting the

sample.

5.3 Results and Discussion

5.3.1 Preparation and characterization of size, PDI and zeta potential

In general, the formation magnetite loaded NPs could be realized by single

emulsion and nanoprecipitation [169,185] . Therefore, these methods have been


- 58 -

chosen to prepare the formulation of PLGA NPs loading magnetite for photoacoustic

imaging and drug delivery [185,187].

Applying the two different preparation methods, particles with clearly different

properties were obtained (Tab.3). By using nanoprecipitation, the resulting NPs

showed a broad size distribution (PDI = 0.53 ± 0.0323) even though the mean

diameter was around 100 nm. As expected, using PLGA, the particles have negative

zeta potential (-8.01 ± 0.81) mV [169].

Table 3. Hydrodynamic mean size (nm), PDI and Zeta Potential (mV) of PLGA NPs loading Magnetite were prepared by nanoprecipitation and single emulsion method

Standard deviation (SD) of (n=3)

On the other hand by using the same concentration of magnetite, single

emulsion resulted in NP with good properties in terms of size and PDI. The size with

250 nm of the particles was much larger than from the nanoprecipitation. The

particles were mostly monodispersly distributed as can be seen (Tab.3) from the

small PDI below 0.1 (PDI = 0.060). From this part, we chose the single emulsion

method for further preparations. The influence of magnetite concentration was then

evaluated for concentrations ranging from 0.5 – 6 mg/mL. Looking at the data

(Tab.4), only the concentrations of magnetite lower than 2 mg/ml resulted in

nanoparticles with small PDI values (monodisperse). High concentrations (4-6

mg/mL) resulted in polydisperse NPs (PDI>0.2). Based on this result, it was

concluded that single emulsion method using magnetite up to 2 mg/mL is a well

defined preparation which could be applied for the main work.

Table 4. Hydrodynamic mean size (nm), PDI and zeta potential (mV) of NPs loading magnetite prepared by single emulsion method.

Concentration of magnetite

Mean Size (nm ± SD)

Polydispersity Index (PDI ± SD)

Zeta Potential (mV ± SD)

0.5 mg/mL 212.18 ± 2.16 0.08 ± 0.02 -5.91 ± 0.35 1 mg/mL 254.38 ± 3.88 0.06 ± 0.03 -5.44 ± 0.42 2 mg/mL 336.29 ± 6.21 0.12 ± 0.04 -5.34 ± 0.61 4 mg/mL 446.29 ± 8.28 0.24 ± 0.06 -4.06 ± 0.72 6 mg/mL 1174.3 ± 12.68 0.54 ± 0.08 -3.51 ± 0.75

Standard deviation (SD) of (n=3)

Methods Mean Size (nm ± SD)

Polydispersity Index (PDI ± SD)

Zeta Potential (mV ± SD)

Nanoprecipitation 115.33 ± 5.01 0.53 ± 0.032 -8.01 ± 0.81

Single Emulsion 254.38 ± 3.88 0.06 ± 0.027 -5.44 ± 0.42


- 59 -

Solvent extraction method which removed the dimethyl formamide (DMF) of

magnetite stabilized with aminohexanamine, could produce particles with good size

and PDI. In the presence of DMF, to mix magnetite with ethyl acetate needs several

steps such vortex and sonication. Therefore for the following preparation of NPs,

DMF was removed. For comparison, oleic acid stabilized magnetite in toluene which

is miscible in ethyl acetate was also utilized. Single emulsion method and

concentrations of magnetite up to 2 mg/mL were applied (based on the previous

result).

The monodisperse magnetite-PLGA NPs were obtained by using 0.5, 1.0 and 2.0

mg/mL (Fig.25) of two kind of magnetite (amino hexanamine-stabilized magnetite and

oleic acid-stabilized magnetite). With a low concentration of magnetite (0.5 mg/mL)

the mean size of particles was around 200 nm, the PDI values indicating a narrow

distribution and the zeta potentials were slightly negative (for both kinds of magnetite

used). Using 1 mg/mL magnetite could also result in monodisperse particles. These

particles have mean size around 220 nm with narrow distributions (PDI 0.04) and

they have also slightly negative zeta potentials (-5.616 mV). This method was

obviously suitable for the preparation of PLGA NPs loading both oleic acid-stabilized

magnetite in toluene (commercially available magnetite from Sigma Aldrich) and

0

50

100

150

200

250

300

1 2 3 4

Size (nm)

-7

-6

-5

-4

-3

-2

-1

Zeta Potential (mV)

Magnetite-aminohexaneamine Magnetite-oleic acid

Samples : A without magnetite B.With magnetite 0.5mg/mL C.With magnetite 1 mg/mL D.With magnetite 2 mg/mL

PDI 0.08

PDI 0.04

PDI 0.03

PDI 0.04

PDI 0.04

PDI 0.09

PDI 0.05

PDI 0.04

Without magnetite A B C D

Figure 25. Size, PDI and Zeta Potential of PLGA NP loading magnetite


- 60 -

aminohexanamine-stabilized magnetite (from Department of Physical Chemistry,

Saarland University). The physical properties of these magnetite-PLGA NPs are

described in Fig.25. According to Fig.25, monodisperse NPs were obtained with

magnetite of 1 mg/mL by using single emulsion method. Therefore, these NPs were

utilized for microscopic characterization and hair shafts imaging.

5.3.2 Imaging NPs by SEM, AFM and TEM Based on the result of the zetasizer measurements, all images were taken

from NPs which were prepared by 1 mg/mL magnetite using single emulsion method.

SEM and AFM measurements in order to know the morphology of the NPs which

were diluted in demineralized water.

The SEM images (Fig.26) reveal that the NPs were spherical and the mean

size was around 176 ± 6.92 nm calculated by freeware ImageJ (Version 1.46). The

AFM image shown in Fig.27 allows to determine the mean size of NPs around 192 ±

17.74 nm by section analysis. Both sizes fit to the size range of particles applicable to

hair follicles. It was also indicating that these sizes were smaller than the

hydrodynamic size. This might be a result of the drying process during sample

preparation for AFM and SEM measurements.

Figure 26. SEM images of the magnetite-PLGA NP

Both SEM and AFM images showed spherical particles well distributed with respect

to their size. As depicted in Fig.27, NP was analyzed regarding their size by marking

the bottom and top of the pancake-like structure by using section analysis. According

to this image s was assumed that the distance between the two arrows indicated the

high of collapsed particle containing a magnetite core.


- 61 -

Figure 27. AFM images of the PLGA NP containing magnetite core

The height was measured to be ~31 nm being a bit larger than the diameter of

pure magnetite NP of ~20 nm (Fig.30). The differences in the value are due to the

polymer surrounding the magnetite particles. It can also be seen that the collapsed

particle is composed out of a thing ring (polymer) and a higher center. From this

structure it can be concluded that the PLGA NPs contain a solid core, the magnetite

particles.

5.3.3 Determination of the successful loading of magnetite in PLGA NPs

Even though we got some indication of the magnetite being incorporated in the

PLGA particles, the key question is still to ensure if the magnetite was successfully

entrapped. To answer the question, TEM, elemental analysis and AFM phase

imaging were applied. TEM images in Fig.28 demonstrate that the grey spherical

PLGA NPs (low contrast) contain black spots (high contrast) being most likely

magnetite (Fig.28A).

The dense particle of magnetite demonstrated a low electron transmittance

which corresponded to the black spot. In comparison, PLGA as organic polymer has

high transmittance therefore the NP was seen as the grey particle. Three analyzed

particles containing magnetite in Fig.28A have the size of approx. 155, 225 and 125

nm which were analyzed by ImageJ.

core

Vertical distance 31 nm


- 62 -

Figure 28. TEM images of the PLGA NPs with magnetite (A), elemental analysis of PLGA NPs loading magnetite (B) and the elemental signature of the TEM grid (C)

To identify the iron oxide, elemental analysis was conducted on one particle

loading magnetite. The NP is a single particle and quite far from another NP therefore

free space could be used as control and the information based on this data that iron

peaks has come from PLGA NP loading magnetite. The strong iron signal from PLGA

NP loading magnetite came from magnetite core (Fig.28B) and the TEM grids did not

have the spectra of iron. The copper and chrome signal were originating from the

TEM grids (Fig.28C).

For comparison with TEM, the phase images of AFM were taken and analyzed

(Fig.29). AFM phase imaging (Fig.29B) refers to record the phase shift signal of tip

oscillation during tapping measurement. The phase shift presents the delay in

oscillation of the cantilever because of different material (adhesion and stiffness). The

images indicate that PLGA NPs contain smaller objects. Together with the TEM

images and the data of the elemental analysis that the observed black spots loaded

in PLGA NPs are magnetite (Fig.29A), these findings (TEM image and phase image

of AFM), could be summed that NPs were formed by two kinds of different

substances [188,189]. Based on the data of TEM and elemental analysis of iron

core

TEM grids

Elemental analysis

A

B

C


- 63 -

signal (Fig.28 A,B,C) also the AFM phase imaging (Fig.29B) it was concluded that

magnetite was successfully loaded into the PLGA particles.

Figure 29. Determination of magnetite core on PLGA NP by TEM and AFM

Figure 30. TEM image of magnetite NPs

5.3.4 Imaging model hair shaft by using CLSM

Imaging of entire/intact hair shafts has been not done yet. The limitation is

regarding to the size and orientation of the hair shaft. Before going to the complex

system such hair shaft in an excised human skin, we tried to create a simple and

appropriate model of a hair shaft in terms of the length and orientation. The model

was created based on agarose (subtitle 5.2.7). In this work we also performed

confocal imaging as control. This measurement was conducted on several model hair

shafts which were filled by fluorescently labeled PLGA NPs applied as cubic sliced

agarose gel blocks. For these experiments different orientations of the model hair

shafts were used as illustrated in Fig.31. It is important in order to obtain the good

images of the model hair shaft because of limitations regarding the focal distance of

the objective.

TEM Image AFM Phase Imaging

A B


- 64 -

Figure 31. Longitudinal and lateral orientation of model hair shaft containing fluorescein PLGA in agarose cubic for CLSM measurement.

In Fig.32 is shown the longitudinal position of model hair shaft. It is also clearly

evidenced that visualizing the model from the top to bottom was not possible. This is

because of the limitation of penetration depth of light and the restricted working

distance of the microscope.

Inside of the model hair shaft, the NPs could not be seen clearly because the void

was covered by one layer of agarose when the top of model was faced objective

lens. The diameter of void is ~0.5 mm (Fig.32) in the range of diameter of human hair

follicles 0.1 - 0.5 mm [190,191] . Only for the lateral orientation the model hair shaft

could be imaged in parts. The composed images are shown in Fig.33. The

distribution of NPs in the model was not homogeneous. Some of NPs were

agglomerated as seen as large and bright fluorescent spots. However, it could not be

fully compared to the hair shaft in terms of the orientation and length not being fully

representative to the real situation. Furthermore, the imaging using several single

CLSM images to compose the structure of interest is time consuming.

Lateral

Longitudinal

Agarose gel

Top

Inside

Top

NPs

Figure 32. CLSM images of longitudinally aligned model hair shaft containing fluoresceinamine-PLGA NP


- 65 -

Figure 33. CLSM images lateral position of model hair shaft containing fluoresceinamine-PLGA NP

5.3.5 Imaging model hair shafts by using photoacoustic microscopy

To answer the difficulties of imaging model hair shaft we introduced a

promising method to image model hair shafts: photoacoustic microscopy. The

measurement followed the procedure described before (section 5.2.8).

Because of the high absorption of magnetite allowing to produce ultrasound

when the laser light was applied, the samples should allow for successful imaging.

The produced ultrasound was transferred as an image by using ultrasound

transducer and the model hair shaft could be visualized.

Therefore, pure magnetite particles (Fig.34C) and PLGA NPs loading

magnetite (Fig.34B) were filled in the model as control blank PLGA NP were used

(Fig.34A). The photoacoustic images are shown in Fig.34. The measurement was

quite fast and relatively easy because taking the image of the different model hair

shafts from the top could be conducted simultaneously. Imaging reconstruction also

allows displaying the side view of model hair shaft as showed in Fig.35.

0 700 1500 µm

NPs


- 66 -

C

B

1 mm

Figure 34. Top view of photoacoustic image of three model hair shafts containing particles. A) pure PLGA particles for control B) magnetite-PLGA NP and C) pure magnetite particles

The tilted orientation is a common position with an angle around 45 to 60° for

hair shafts [192,193]. The size and the orientation did not cause any problems in

imaging. Based on Fig.35, the length of the model hair shaft is around 1.1 mm.

Due to the length distribution of human hair follicles in the range of 0.5-3 mm

[194], we assumed that 1.1 mm is a valid size for our model being at least in the right

range. Additionally, it was seen, that the covering layer made of agarose does not

limit the imaging process in terms of the depth and the size of model hair shaft.

Oberfläche

Probe C

Figure 35. Photoacoustic images representing a side view of a model hair shaft containing magnetite-PLGA NP (red ellipse), the dashed line indicates the location of the covering layer.

A

Covering layer Surface


- 67 -

5. 4. Conclusions

In the present investigation, the formulation of PLGA NPs loading magnetite

was obtained. The NPs were prepared by nanoprecipitation and single emulsion

methods. By using single emulsion method in contrast to nanoprecipitation, spherical

particles could be produced with a narrow distribution. By applying higher

concentrations of magnetite (≥2 mg/mL) the NPs have shown broad unwanted

distribution of particles. The successful incorporation of magnetite into the polymer

NPs could be determined by using AFM and TEM images in combination elemental

analysis. PLGA NPs loading magnetite were then applied to visualize model hair

shafts embedded in agarose gel representing the hair follicle dimensions. The model

hair shafts containing PLGA NP loading magnetite could be easily imaged by

photoacoustic microscopy. Hence photoacoustic microscopy is an appropriate and

promising technique to visualize intact hair follicles as large objects. This technique

could be used to overcome the limitations of classical CLSM technique regarding the

large scale and orientation of hair follicles for determining the localization of drug

carrier systems.

- 68 -

VI. SUMMARY

Hair loss is a case which can be found in men and women. Usually, if hair loss

occurs in an every day manner, it will grow again by regeneration of new hair. When the

amount of hair loss is more than the amount of hair growth, it will decrease the total

amount of hairs. This condition could happen everyday and cause baldness. Feared by

everyone, baldness is caused by continuous hair loss. Disruption of hair growth is

influenced by internal and external factors of human body. Baldness is a rare case.

Although rare, this is a worrying case for the patients. Nowadays, it has been attempted

the way to prevent and treat hair loss by commercial products such as lotions, creams

and shampoos. These commercial products containing caffeine in non particulate

formulations are available on the market. Based on the recent research, it was known

that the particulate formulation has better penetration into the hair follicles. A particulate

formulation still needs to be developed, especially a nanoparticulate formulation which is

promising product for hair follicles application. The investigations of this kind of

formulation have been indicating the significant progress. Some of these investigations

are aimed to develop pharmaceutical dosage forms because only a limited number of

them employed nanoparticles (NPs). In the pharmaceutical field, NPs are produced with

a uniform size. This NPs loading drug are intended to deliver and target the drug to the

human body. Because NPs can reach the specific target, this should be able to diminish

the side effect of drug.

Based on these informations, this study aimed to develop and characterize a

particulate formulation by utilizing a biocompatible and biodegradable polymer such as

chitosan and poly(lactic-co-glicolic acid) (PLGA) to form NPs loading caffeine.

Interaction between chitosan and caffeine was determined by Fourier Transform Infrared

Spectroscopy (FTIR), Differential Scanning Calorimetry (DSC) and by a solubilization

study of caffeine. FTIR and DSC measurements indicated that the interaction between

chitosan and caffeine is based on the formation of a complex. The solubilization study

- 69 -

showed the ability of chitosan to solve insoluble caffeine in a saturated dispersion and

increase the saturation concentration. NPs loading caffeine were prepared by combining

chitosan and PLGA to form chitosan-PLGA NPs (by single emulsion method) and solely

PLGA NPs (by double emulsion method). Data of NPs characterization showed that NPs

have a spherical shape. Especially, NPs made from chitosan-PLGA showed better

properties than that of PLGA NPs in terms of particle size and size distribution. Loading

of caffeine was increased by using chitosan. Caffeine release from NPs is slower than

passive diffusion of solved caffeine which is not formulated in the form of particles.

The difficulty of delivering drug across the skin is prevalent. One of the strategies

gaining more attention over the last years is the targeting of the hair follicles. Upon

application the carrier system accumulates in the hair follicles and can release its load.

The common/standard visualization technique is confocal laser scanning microscopy,

but it has several problems originating from the physical size of the hair follicles and their

orientation in skin. An intriguing alternative to fluorescence-based techniques is

photoacoustic microscopy. Optoacoustic imaging is a novel method that based on the

specific absorption of light in the near infrared region. The absorber such as iron oxide is

irradiated by short laser pulses in the sub-nanosecond range generating broadband

acoustic signals as a result of the thermoelastic effect. These signals propagating

through the samples can be obtained with adequate ultrasound transducers. When

compared with conventional ultrasound imaging, optoacoustic techniques have the

advantage of higher contrast. Optoacoustic signal amplitudes are proportional to the

local absorption coefficient µa. Since the absorptions coefficient is dependent on the

concentration of absorbing material, this effect can be utilized in order to assess the

suitability of different material types (magnetite or magnetite-loaded polymer) as contrast

agents.

As a possible label a magnetite core was introduced to the polymeric PLGA

particles synthesized for drug delivery purposes. This combination of materials allows

achieving magnetic properties as well as biocompatibility and biodegradability of the

carrier. Furthermore, a good model system mimicking hair follicles would facilitate ex

vivo investigations. Hence, preparation and characterization of magnetite PLGA NPs

was in focus and the evaluation of a model hair shaft which could be imaged by

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photoacoustic microscopy. The model hair shaft was prepared by using a syringe which

was immersed into a 1.75% agarose gel (in a Petri dish). After solidification and

retraction of the syringe, a void had been formed (serving as the model). This void was

then filled with a suspension of NPs. The results showed that magnetite-PLGA NPs

were successfully produced by utilizing 1 mg/mL magnetite and the model hair shaft

containing magnetite-PLGA could consequently be imaged by using photoacoustic

microscopy while confocal microscopy imaging was time consuming and complicated

which could not image the large structure.

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ABBREVIATIONS

AFM = Atomic force microscopy

API = Active pharmaceutical ingredient

AuNPs = Gold nanoparticles

cAMP = Cyclic adenosine monophosphate

Chit = Chitosan

CLSM = Confocal laser scanning microscopy

Da = Dalton

DHT = Dihydrotestosterone

DLS = Dynamic light scattering

DMF = Dimethyl formamide

DMSO = Dimethyl Sulfoxide

DSC = Differential scanning calorimetry

EE = Encapsulation efficiency

FDA = Food and drug administration

FDC = Franz diffusion cell

FTIR = Fourier transform infrared

FWHM = Full width half maximum

HF = Hair follicles

HPLC = High performance liquid chromatography

MNPs = Magnetite nanoparticles

MRI = Magnetic resonance imaging

MT = Magnetite

MWCO= Molecular weight cut off

NPs = Nanoparticles

PAM = Photoacoustic Microscopy

PBS = Phosphate buffered saline

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PDI = Polydispersity index

PLGA = Poly(lactic-co-glycolic acid)

PVA = Polyvinyl alcohol

SLNs = Solid lipid nanoparticles

SC = Stratum corneum

SEM = Scanning electron microscopy

5α−Red = 5α−Reductase

TEM = Transmission electron microscopy

TPP = Tripolyphosphate

UV = Ultraviolet

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Curriculum Vitae Name : Mardiyanto

Date of birth : 10 March 1971

Place of birth : Padang, West Sumatra. Indonesia

Nationality : Indonesian

Education: 1999-2002: Master of Science (MSc) at the Department of Pharmacy, Faculty of Natural Science, Bandung Institute of Technology (ITB Bandung), Jalan Ganesha 10, Bandung, West Java, Indonesia. Thesis was entitled: "Enhancement the tetracycline production of Streptomyces aureufaciens". 1995-1996: Apoteker (Pharmacist) at Department of Pharmacy (professional program of pharmacist) Faculty of Natural Science, University of Andalas, Kampus Unand Limau Manis, Padang, Indonesia. 1990-1995: Bachelor of Science (BSc) at Department of Pharmacy, Faculty of Natural Science. University of Andalas, Kampus Unand Limau Manis, Padang, Indonesia. Best graduate from Faculty of Natural Science, University of Andalas, Padang, Indonesia 1990: Completed high school education at the Indonesian state high school SMAN 3, Jalan Gajah Mada, Padang, Indonesia. Career Summary: 2009- Present: PhD student at Department of Pharmaceutical Nanotechnology, Saarland University, Saarbrücken, Germany. 2007-2008: Training on International Management Competence of Industrial Biotechnology, Germany.

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2003- 2007: Indonesian government employer: Assistant lecturer at pharmaceutical study program, Faculty of Natural Science, Sriwijaya University, Inderalaya, Indonesia. 1998- 2000: Pharmacist at Apotek Bunda, Palembang, Indonesia 1997- 1998: Academic facilitator at Harapan Bangsa, Pharmaceutical College Palembang, Indonesia Publications: Research articles:

� Mardiyanto and Marc Schneider, "Investigation of Nanoparticulated Formulation Loading Caffeine” J. Drug Delivery Letters, 2013. In Process of Submission

� Mardiyanto, Clemens Tscheka, Wolfgang Bost, Marc Fournelle and Marc Schneider, “Photoacoustic Imaging of Model Hair Follicles Containing Magnetite-PLGA Nanoparticles”. 2013. In process of submission

� Mardiyanto and Budi Untari, Chitinase of bacteria from Inderalaya swarm, J. Kedokteran UNSRI Palembang Indonesia. 2005, 216: p.34-39.

Poster presentations:

� Mardiyanto, Clemens Tscheka, Marc Fournelle and Marc Schneider, “Photoacoustic Imaging of Model Hair Shaft” Inascon Conference. August. 2012.

� Mardiyanto and Marc Schneider, “Preparation and Characterization Chitosan coated PLGA NPs Loading Caffeine”, 8th international conference and workshop on Biological Barriers- in vitro tools, nanotoxicology, and nanomedicine, Saarland University, Saarbrücken, 21st March – 1st April 2010. Other Scientific Activities: � Participation in the association of the Pharmacist in South Sumatera Indonesia, development of nature resources for traditional drugs supplying, 2003-2007. � Participation in the Dikti development of research collaboration inter University in Indonesia. 2006-2007.

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Grants: � Indonesia Dikti Scholarship for a 3 years study at Saarland University for a PhD degree 2009-2012.

� International management competence on Industrial biotechnology, one year in Germany, June 2007-May 2008.

� Indonesia Dikti Scholarship for a 2 years study at Bandung Institute of Technology (ITB) Bandung, Indonesia for a MSc Degree 1999-2001.

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Acknowledgments

Above all, I am really thankful to Allah for his infinite blessings of my

destiny. Thanks Allah the most merciful for accepting my prayers, protecting

me and keeping me on track during the life. It is a pleasure time to convey my

gratitude to all of persons though I can only name a fraction of them in my

honest acknowledgments:

First of all, I would like to express my many sincere gratitude and

appreciation to my supervisor Prof. Dr. Marc Schneider for offering me the

opportunity to join his research group. I remembered first time I came to him at

July 2007 for my purpose to learn much about nanoparticles. His spirit came to

me to win the Grant “Indonesia Dikti scholarship” and finally I met him again

at October 2009 as PhD student. His support and encouragement was spending

a lot of effort and time on this study, nice discussions especially regarding the

new ideas how to solve the problems in our investigation. I also want to thank

him for being understanding and guiding me for that. Far away from my

parents, sister and brother in Indonesia, I feel have family in Germany. I am

really thankful for everything.

I would like also to thank my entire research group: Achim Nauert

introduced me the laboratory system in our institute. Thank you very much is

attributed to his sharing experience of preparation of nanoparticles and AFM

measurement. I would like say thanks to Clemens Tscheka for his friendship

and capability to encourage my knowledge about SEM and TEM. Dr. Nico

Reum, Dr Hagar Ibrahim, Qiong Lian, Dr Ke Li, Dr Xavier Le Guével, Saeed

Ahmad Khan, Daniel Primavessy, Dr Noha Nafee, Afra Torge, René Rietscher,

Carolin Thum, Nazande Ghünday, Asli Katkilic, Kristina Malinovskaja, Shi

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Chen and Mohamed Tawfik for always acting like a team and helping in

numerous ways.

I am grateful to Prof. Dr. Ulrich Schäfer for sharing with me his attention

and motivation for me. His valuable guidance and suggestions has often led

me to the right scientific direction.

Thanks are extended to Prof. Dr. Claus-Michael Lehr for his support,

motivation and encouragement with nice way to develop a lot a new finding in

research. Thank you very much to Dr. Nicole Daum was sharing with me her

experience in safety work in laboratory, to establish the condutcive daily

activities in laboratories. Thank you attribute to Dr Brigitta Loretz for scientific

discussion and nice working in Laboratory.

Thank you also attribute to Dr. Tsambika Hahn. She kindly spent time

and effort to show me some of the valuable Franz Diffusion Cell and sampling

techniques that I have practiced throughout my PhD work, Dr Christian Ruge

for nice friendship and helping to prepare magnetite NPs and use several

software to analyze the raw data of zeta sizer, DSC and nanosight, Leon Muijs

for helping me with many things especially for CLSM measurement. Peter

Meiers for his help with HPLC and solve problem about computer and Petra

König for introducing and administrate material for our laboratory. Lutz

Franzen and Christiane Mathes (for help taking image of hair follicle), to both

of you: thank you very much for friendship, discussion and helping regarding

the nice hair follicles. Sarah Müller, Karin Gross and Isabelle Conrad-Kehl,

thank you very much for helping the academic administration.

Thank you very much to Dr. Marc Fournelle and Dr Wolgang Bost for

helping the photo acoustic measurement in IBMT St Ingbert Germany. Their

friendship was helping me for an interesting of photoacoustic measurement.

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I would like to acknowledge the Directorate of Higher Education of

Indonesia for the financial support during my stay in Germany. I would like

also to say thank you very much to my Rector of Sriwijaya University, Dean of

Faculty of Natural Science and Head of Department of Chemistry and

Pharmaceutical Study Program of Faculty of Natural Science, Sriwijaya

University.

A PhD program was not only about a scientific achievement, this was

also a happy time and friendship. In our international institute, I enjoyed

working with kind and helping friend as much as: Salem, Hiroe, Nico Mell,

Christina, Ratnesh and Prajakta also Ankit, Hussein, Babak, Anne and Ana.

For my Mom and Dad, thank you very much for your praying. Your

advice and your patient had already guided me to see many advantages of

knowledge in Germany. You will be in my heart always and forever. My sister

and my brother, thank you very much for your time and spirits. For my niece

Cici, thank you very much for your time and praying. After all the time we will

see how the life should life and how the knowledge can develop the human

life. My sincere thanks to all Indonesian friends in Germany and I would like to

say many thanks for helping and friendship to everybody who I can not say all

of them in this chapter.

Saarbrücken, 4 June 2013

Mardiyanto

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von mardiyanto saarbrücken 2013

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