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Tracking Dermal Drug Delivery

6.  Tracking Dermal Drug Delivery

Stimulated Raman scattering microscopy was used to gauge the permeation of topically applied pharmaceuticals and formulation solvents into pig dermis. This chemically selective technique creates high-resolution 3D images, from which semi-empirical information may be extracted. Ibuprofen, applied as a near-saturation solution in propylene glycol, was directly observed to crystallise in/on the dermis, as the co-excipient permeated more promptly, resulting in precipitation of the drug. Coherent Raman scattering microscopy is also an excellent tool, in conjunction with more conventional confocal fluorescence microscopy, with which to image micro/nanoparticle-based formulations. Specifically, the uptake of particles into thermal ablation transport pathways in the dermis has been examined.

6.1.   Introduction

The dermis, as described in chapter 2, is an effective barrier, which has evolved to prevent entry of undesirable foreign substances into the body. However, despite the associated difficulties, topical drug delivery remains an attractive therapeutic approach (especially for the treatment of dermatological diseases) since it avoids the need for systemic exposure and its related disadvantages. To deliver pharmaceuticals both safely and effectively through the dermis, it is necessary to quantify the penetration of the active agent and to understand the evolution of the drug vehicle upon application and the disposition of key components of the vehicle. Currently, a widely-used approach involves removal of the stratum corneum (SC) by sequential adhesive tape stripping at specific time points, post-application of a formulation [1]. This enables generation of concentration profiles of the active and certain formulation ingredients; however, this method is both invasive, painful, and labour-intensive. In addition, it does not reveal information such as the pathway taken by the drug or the reasons for the poor penetration that is typically observed.

Coherent Raman scattering (CRS) microscopy provides a new tool with which to tackle this problem. CRS as a label-free imaging technique is capable of real-time, non-invasive examination of living cells and organisms based on molecular vibrational spectroscopy. A coherent non-linear Raman signal is created by focussing two synchronised ultrafast pulse trains into a sample with a difference in frequency matched to a Raman active mode of a molecular species of interest. The non-linear nature of this process confines the signal to a sub-micron focal volume that can be scanned in space, allowing three-dimensional mapping of bio-molecules in tissue with sub-cellular resolution.

CRS microscopy may be achieved by detecting either coherent anti-Stokes Raman scattering (CARS) [2, 3] or stimulated Raman scattering (SRS) [4-10]. In CARS, the coherent anti-Stokes signal created at frequency ωas = 2ωp − ωS, is spectrally isolated from the pump and Stokes beams using filters, and a signal intensity map can be obtained by raster scanning the pump and Stokes beams across the sample, allowing visualisation of the location of biomolecules of interest. The chemical specificity, strong signal strength and non-invasive imaging capabilities offered by CARS have been exploited in a wide range of biological studies, providing diverse information such as: the spectroscopic properties of diseased and healthy dermis samples [3] and the interaction and uptake of nanoparticles within organs such as the liver [11] and the brain [12], as well as in living cells [13]. In contrast to CARS, SRS relies on detecting subtle changes in the intensities of the excitation fields that occur by virtue of stimulated excitation. When the difference frequency, ωp − ωS, matches a molecular vibrational frequency, the intensity of the Stokes beam, IS, experiences a gain (SRG), ∆IS, while the intensity of the pump beam, Ip, experiences a loss (SRL), ∆Ip. The intensity transfer from the pump to the Stokes beam only occurs when both beams are incident upon the sample and can be detected with high sensitivity using modulation transfer detection. Modulating the intensity of the Stokes beam modulates the SRS process and hence transfers an intensity modulation onto the pump beam. The amplitude of the transferred intensity modulation is directly proportional to the concentration of target molecules and by modulating at frequencies (~1.7 MHz)  above the laser noise, can be detected with a lock-in amplifier with great sensitivity (1 in 106).

For quantitative studies, SRS is often the more appropriate technique because the signal output is identical to the spontaneous Raman spectrum, and its linear concentration dependence affords a relatively straightforward quantitative analysis. These properties have proven to be efficacious in studies investigating drug interactions with dermis both in vitro and in vivo [6, 7]. CARS creates more complex vibrational spectra, including the presence of wavelength-shifted peaks, negative contrast, and a quadratic concentration dependence, making numerical image interpretation much more challenging. However, for less quantitative studies, such as imaging particle-dermis topology, CARS can be an excellent tool, since the high concentrations of chemical species present in particles create strong signals due to the quadratic concentration dependence.

In the first part of this study, we demonstrate the differences in appearance between mouse and pig stratum corneum (SC), the latter representing (as is widely accepted) a more appropriate model for the human barrier. Subsequently, ketoprofen is applied to pig dermis as a solution in perdeuterated propylene glycol. The architecture of the dermis is imaged based on the contrast of the CH2 groups present most notably in the dermis lipids. The disposition of ketoprofen is visualised using the aromatic CC contrast, and the propylene glycol-d8 by the CD bond. The data are compared to previous experiments on mouse ear dermis [6].

We also demonstrate the suitability of these imaging techniques for characterising the effects of a device designed to aid the (trans)dermal delivery of compounds which permeate the dermis particularly poorly; for example, macromolecules or very water-soluble and/or highly charged compounds. Thermal ablation creates small pores or channels within the SC by application of a short burst of heat, in this case, provided by a microarray of metal filaments. This serves to increase the permeability of the outer layer of the dermis’s barrier, without damaging the deeper layers of tissue. Here, we investigate the suitability of CRS microscopy, in combination with confocal fluorescence microscopy, for characterising the new transport pathways created by such a device, and the disposition of topically applied micro- and nano-particles within them.

6.2.  Materials and methods

Pig abdominal dermis was cleaned with water, trimmed to remove excess hair and dermatomed to a nominal thickness of 300 μm. In an initial set of experiments, the dermis was dosed with a solution of ketoprofen (Sigma-Aldrich, Gillingham, UK) in per-deuterated propylene glycol (Isotec, Ohio, USA), mounted for imaging as described below and then examined by SRS/CARS; the concentration of the drug in propylene glycol was 180 mg/mL. A short parallel study was also conducted in the same way during which the dermis was treated with a solution of deuterated ibuprofen (Sigma-Aldrich, Gillingham, UK) in undeuterated propylene glycol (Sigma-Aldrich, Gillingham, UK) at a concentration of 380 mg/mL, corresponding to approximately 90% saturation [14]. For the ablation experiments, dermis was treated with a poration device (PassPort™ A5A Applicator, Altea Therapeutics, Atlanta, GA, USA) immediately prior to mounting in a vertical Franz diffusion cell (Permegear, Hellertown, PA, USA), exposing a diffusion area of 2 cm2. The receptor chamber was filled with phosphate-buffered saline solution (7.5 mL, pH 7.4). Particle suspensions (500 μL) were applied to the dermis surface exposed in the donor chamber, which was subsequently covered with Parafilm® to prevent evaporation. The sample was incubated at 37 °C for a chosen duration, after which the Franz cell was dismantled, the particle suspension removed, and the dermis immediately mounted for imaging.

Yellow–green fluorescent 20 nm and 2 μm diameter carboxylate-modified FluoSpheres® were purchased from Invitrogen (Eugene, OR, USA). Nanoparticles were prepared by free radical polymerization under a nitrogen atmosphere for 3 h from: ultrapure water (50 mL), sodium dodecyl sulphate (0.25 g, Sigma-Aldrich, Gillingham, UK) and methyl methacrylate (3.25 g, Sigma-Aldrich, Gillingham, UK). The resulting mixture was heated to 75 °C, and the reaction was initiated with potassium persulphate (25 mg, Sigma-Aldrich, Gillingham, UK). For deuterated particles, methyl methacrylate-d8 (Sigma-Aldrich, Gillingham, UK) was substituted, and for the fluorescently labelled particles, fluorescein O-methacrylate (32.5 mg, Sigma-Aldrich, Gillingham, UK) was mixed with the methyl methacrylate before addition. The average particle diameter of 40 nm (with a polydispersity index of 0.1) was determined using dynamic light scattering (Zetasizer Nano S, Malvern Instruments Ltd, Worcestershire, UK).

All samples were mounted for imaging between two glass coverslips (22 × 50 mm) cover glasses (Menzel-Gläser, Braunschweig, Germany). The slides were sealed together to reduce the loss of volatile substances and to minimise swelling/dehydration of tissue; the latter is of particular importance for lengthy time-course experiments. This was achieved using rectangular frames constructed from Parafilm. The frames were placed onto glass coverslips and the back of the slide was warmed on a heated stirrer-plate at 60 °C for a few seconds until the Parafilm became translucent. After cooling, the sample was placed within the confines of the frame, and a top coverslip applied. This glass slide was then sealed to the second one using a soldering iron, to melt the Parafilm, which then fused the two slides together. Utmost care was taken to contact the heat source only around the outer edges of the frame to prevent heat from reaching the sample. The glass slides were then secured inside a brass flow cell clamp for imaging. This served to prevent flexing of the thin, pliable glass coverslips during the study. Laser power was minimised throughout to prevent damage to the biological samples examined.

CRS and two photon fluorescence (TPF) imaging were carried out using a custom-built multi-modal microscope comprising of a modified inverted microscope and confocal laser scanner (IX71 and FV300, Olympus UK). To optimise the transmission of the near IR light needed for CARS and SRS imaging, the standard galvanometer scanning mirrors were replaced with silver galvanometric mirrors and the tube lens was replaced by a MgF2 coated lens. The dichroic mirror within the scan unit was replaced by a silver mirror, which gave high reflectivity throughout the visible and NIR (21010 Chroma Technologies, Vermont, USA ). The light was focused onto the sample using either a 60× 1.2NA water immersion objective or a 20× 0.75NA air objective (UPlanS Apo, Olympus UK).

TPF was excited using a 800 nm output from a mode-locked femtosecond Ti:Sapphire laser (Mira 900 D, Coherent) with a pulse width of approximately 100 fs and a 76 MHz repetition rate (Fig. X). The signal was collected in the epi-direction using the objective lens and separated from the laser fundamental using a long pass dichroic mirror (670dcxr Chroma Technologies) and detected by a PMT (R3896, Hamamatsu) with filters (CG-BG-39 and F10-400-5-QBL, CVI laser) to isolate the TPF signal.  Synchronised, dual-wavelength picosecond excitation required for CRS was provided by an optical parametric oscillator (OPO) (Levante Emerald, APE-Berlin) synchronously pumped at 532 nm by a frequency doubled Nd:YVO4 laser (picoTRAIN, High-Q GmB), delivering 7 ps pulses at a repetition rate of 76 MHz. The pump-laser fundamental (1064 nm) was also available as a separate output. The OPO uses a temperature-tuned, non-critically phase matched LBO crystal to allow continuous tuning of the OPO signal from 690 to 980 nm by adjustment of the LBO temperature and an inter-cavity Lyot filter. The OPO signal and pump-laser fundamental were used as the pump and Stokes beams, respectively, for CARS and SRS, and as the probe and excitation beams for two photon photothermal lensing. The two pulse trains were spatially overlapped on a dichroic mirror (1064 DCRB, Chroma Technology) and temporally overlapped using a delay-stage. Before entering the microscope, the 1064 nm beam was amplitude modulated at 1.7 MHz using an acousto-optic modulator (Crystal Technologies Inc., type S/N 3080-197).

To computer

Forward detector (HCARS)



Half-wave plate



1064 nm


Delay stage

532 nm

Epi-detector (epi-CARS)


Pump beam


Scanning box


Figure X: Schematic diagram of the SRS setup.

The CARS signal was detected in the epi-direction by the objective, spectrally isolated from the pump and Stokes beams using a dichroic mirror (750LP, Chroma) and band pass filter (750/210, Chroma) and detected using a red-sensitive PMT (R3896, Hamamatsu) mounted on the rear port of the microscope. SRS was detected in the forward direction by a 1.0NA condenser lens (LUMFI, Olympus) and a large area photodiode (FDS1010, Thorlabs). A bandpass filter (850/90 nm, Chroma Tech) was mounted in front of the detector to block the modulated 1064 nm beam. A lock-in amplifier (SR844, Stanford Research Systems) was used to detect the SRS signal with a time constant of 30–100 μs. SRS images were created by recording either the in-phase ‘X’, or magnitude ‘R’ outputs from the lock-in. Images were processed using ImageJ, a Java-based image processing program [15].

Laser scanning confocal microscopy images were obtained using a 510 Meta inverted confocal laser scanning microscope (Carl Zeiss, Jena, Germany). Samples were excited using a 405 nm (diode) and a 488 nm (argon) laser. EC Plan-Neofluar M27 objectives were used for image acquisition (40×/1.30 oil DIC, or 20×/0.50 air). Fluorescence signals were recorded as separate channels at 420–480 nm (blue), and 505–530 nm (green).

6.3.  Results

  1. Pig vs mouse ear dermis

Images of un-dosed tissue were recorded for both pig dermis and mouse ear dermis ( Fig. 1). The mouse ear dermis created very clear images, in which the hexagonal shaped corneocytes are clearly visible, framed by the strong signal created by the intercellular lipids. Pig SC, however, showed considerably less structural detail. The pig dermis used for this experiment had been specially prepared with a scalpel to ensure comparable thickness to mouse ear dermis, yet individual corneocytes were still difficult to identify. While mouse SC typically has 3–5 columnar-stacked layers of corneocytes, pig SC is composed of approximately 20 layers of these cells, which are less regularly arranged but more densely stacked [16]. It is possible, therefore, that this higher stacking density in pig dermis gives rise to convoluted corneocyte layers within the experimental z-resolution (1 μm) of the technique, thereby obscuring the finer detail visible in mouse SC.

Figure X: SRS images obtained at 2855 cm−1 for CH2 contrast. Left: mouse ear SC. Right: Pig SC.

In this study, SRS was recorded in the forward direction, so samples were chosen to be as thin as possible without compromising the integrity of the barrier. It was found that dermatoming to 300 μm was optimal. It should also be noted that mouse ear dermis was taken from white mice because the melanin pigmentation present in brown mouse dermis absorbs much more strongly than its surroundings, causing localised burning, and making imaging much more challenging. The pig dermis used was therefore screened carefully to avoid pigmentation spots.

Although the mouse ear dermis displayed more prominent features, its barrier function is recognised as being inferior to that of the human counter-part. Pig dermis, which was used here, on the other hand, is widely accepted as an excellent model for the human tissue [17].

Figure X: SRS X–Z orthogonal view images of pig dermis dosed with ketoprofen in propylene glycol-d8. Red/orange: 2855 cm−1 CH2 stretching frequency. Cyan: PG-d8 visualised at 2120 cm−1 (CD2 stretch). Magenta: ketoprofen aromatic ring CC stretch at 1599 cm−1. Scale bar represents 100 μm.

6.3.2.  Drug permeation time-course experiment

A time course experiment on pig dermis dosed with ketoprofen in deuterated propylene glycol was performed by sequentially tuning the lasers between 2855 cm−1 to image the dermis lipids, to 2120 cm−1 to image CD contrast of the propylene glycol, and then to 1599 cm−1 to obtain contrast for the aromatic CC stretch of the ketoprofen. A fourth off-resonance wavelength (1554 cm−1) was also recorded to verify that the signal vanished when the wavelength was tuned away from the resonance frequency.

It was subsequently possible to extract quantitative information from the series of images (XZ orthogonal views of the 3D image stack are displayed in  Fig. 2). However, for superior analysis, a number of factors needed to be carefully addressed: signal loss with depth, due to scattering by the dermis, fluctuations in laser intensity, sample movement/swelling, and sufficient formulation coverage to assume an excess for the experimental duration.

The average pixel intensity of each of 5 delimited regions (20 by 20 pixels) was determined using ‘specify’ and then ‘plot z-stack profile’ plugins in ImageJ to create profiles of the SRS signal versus depth into the dermis.  Regions for analysis were selected on the basis of their sufficiently thick coverage of formulation. SRS signal was normalised against the signal recorded at a secondary detector (using a beam splitter) to account for any fluctuations in laser intensity. The surface of the dermis within each region was defined by taking the first derivative with depth of the SRS signal at the CH2 frequency, and designating the maxima as ‘0 μm’ sample depth. It should be noted that the surface of the dermis is quite uneven; therefore ‘0 μm’ should be considered an approximation. Adjustments were made where necessary to correct for small movement shifts, detected by SRS z-stacks taken periodically. The SRS signal (average pixel intensity) at the surface ‘0 μm’ on the first time point was designated 1.00 and subsequent depths and time points were plotted as a fraction of this value. Small amounts of background signal (e.g., from particles of dirt), detected at the off-resonance wavelength were separated from the SRS resonant signal before normalisation.



Figure X: Normalised SRS signal intensity as a function of depth into the dermis for: a) Propylene glycol-d8, and b) Ketoprofen. Average pixel intensity was determined using the ‘plot z-axis profile’ function in ImageJ.

 Fig. X shows the semi-quantitative data extracted from a series of images recorded over 2 h. The profiles show similar features to those obtained for ibuprofen using the tape stripping approach and human dermis [14, 18]. Notably, permeation occurred more slowly than determined previously by SRS for mouse dermis [6], as would be expected. A visual inspection of the orthogonal images of the ketoprofen signal reveals that, over the duration of the experiment, the drug began to crystallise out and form a solid layer on the surface of the dermis. Close inspection shows a uniform intensity band of ketoprofen on the surface of the dermis after 120 min.

An even more extreme example of this phenomenon was observed with ibuprofen, as had been previously reported using mouse dermis [6]. Following application of a propylene glycol solution of deuterated ibuprofen-d3 to the dermis at a concentration close to its saturation solubility, large, geometric crystals were clearly visible, having formed with-in 30 min post-application of the formulation ( Fig. X).

Figure X: Crystals of Ibuprofen-d3 formed in/on the dermis within 30 min post application. Panel (a): SRS contrast was obtained at 2120 cm−1 corresponding to CD stretch. Panel (b) depicts a still from a video rotation of a 3D projection created by ‘colour merge’ of the CD contrast (blue, for ibuprofen) with the corresponding CH2 signal emanating from the dermis lipids (red) to reveal the topology.

6.3.3. Particle uptake into thermal ablation channels

Confocal fluorescence microscopy was first used to image yellow– green fluorescent, 2 μm diameter microparticles (FluoSpheres®) on dermis, which had been pre-treated with a thermal ablation device. In the images presented below, the green signal at 488 nm corresponds to the fluorescein label on the FluoSpheres®, while the blue emission at 405 nm reflects dermis autofluorescence.

In panel (a) of  Fig. X, the confocal microscopy image shown was obtained using a ‘tile’ function, to stitch together a 5 × 5 field-of-view grid to enable visualisation of the array of rectangular, trench-like structures created by the device. Panel (b) focuses on a single ‘trench’ created by the porator, and individual particles are clearly visualised. Panel (c) displays the corresponding orthogonal view of this image stack. Dermis auto-fluorescence is heightened around the trench boundaries, which may be caused by tissue compaction, or localised burning post-ablation. Panel (d) plots the variation in fluorescence across the orthogonal image at the surface, mid-depth and base of the channel, corresponding to the fluorescent particles adhering to the topology of the structure.


Figure 5:  Confocal microscopy images of 0.02 μm (panel a) and 2 μm (panels b and c) FluoSpheres® (green, 488 nm) on thermally ablated pig dermis (visualised via its auto-fluorescence (blue) at 405 nm). Panel (a): Tile view stitched images to reveal the patterned array of the ‘trenches’ created by the device. Panel (b): Projection from above of an individual ‘trench’. Panel (c): Orthogonal view of the panel (b) image stack. Panel (d): Fluorescent signal intensity profiles transecting the channel at the surface, mid-depth, and base (average values plotted across 10 pixels).

The images confirm that the channels created by the device are approximately 300 μm long, 50 μm wide, and up to 100 μm deep (z-direction). The 3D images obtained reveal the clustering of particles on the surface of the dermis, within natural creases, and within the channels created by the device. No evidence was observed for the permeation of FluoSpheres® outside the boundaries of the pores.

In addition to confocal microscopy, the experiment was also performed using a combination of TPF and CARS to reveal further detail of the 3D architecture of the dermis ( Fig. 6). Panel (a) of  Fig. 6 illustrates a thermally porated ‘trench’ in the dermis, where the CARS signal (red) from endogenous lipids is absent and replaced only by a dark rectangle. This surface image reveals the presence of the green–yellow microparticles in the ‘crevasses’ of the dermis and at the edges of the trench, in a manner consistent with that observed in  Fig. 5.

Figure 6: Porated dermis incubated for 1 h with 2 μm diameter FluoSpheres®. CARS contrast (CH2 stretching) was recorded at 2855 cm−1 (red). TPF was recorded at 813 nm (green). Images were recorded at 1 μm depth increments. Panel (a): XY view of a channel. Panel (b): 3D cross-section of the channel (prepared using the ImageJ ‘volume viewer’ plugin). Panel (c): Fluorescence signal intensity profiles transecting the channel at the mid-depth and base.

Panel (b) of  Fig. 6 shows a 3D reconstruction of a ‘trench’ with microparticles visible along the walls and at the base of the pore created by the device. The fluorescence signal intensity profiles in panel (c) provide a semi-quantitative confirmation of this visual observation. At the mid-depth of the ‘trench’, a substantial signal from particles that adhered to the walls is detected while there is no detectable fluorescence from the (presumably empty) centre of the pore. In contrast, at the base of the trench, as might be expected, there is accumulation of particles and a large fluorescent signal.

Although fluorescence-based imaging techniques are suitable when studying large particles, which can be individually visualised, smaller-sized particles present a greater challenge for accurate analysis. The subsequent experiments, performed using nanoparticles of 40 nm diameter that were prepared from a methyl methacrylate monomer, incorporating a small amount of fluorescein to enable visualisation using TPF, illustrate this point.

Panels (a–c) of  Fig. 7 show fluorescently labelled nanoparticles on the dermis surrounding an ablation pore. Since the dermis itself produces auto-fluorescence, it is difficult to determine whether the low-level signal is originating from a small amount of particles or from the dermis itself, because the particles are too small to be individually distinguished. For example, panel (b) shows a confocal slice mid-depth through the trench; while it was expected (and it is most probably the case) that the nanoparticles would be confined to the pore, there is a confounding green haze to the surrounding tissue. The cross-section in panel (c), while apparently confirming the anticipated distribution of particles to the walls and base of the ‘trench’, does not entirely remove this potential ambiguity.

Figure 7: Porated dermis incubated for 12 h with 40 nm diameter particles. CARS contrast was re-corded at 2855 cm−1 (CH2, red). Panels (a–c): Fluorescent methyl methacrylate particles; TPF was recorded at 815 nm (green); images were recorded at 0.5 μm depth increments. Panels (d–e): Deuterated methyl methacrylate particles; SRS (CD2) at 2120 cm−1 (blue); images were recorded at 1 μm depth increments.

Consequently, a second batch of nanoparticles was prepared; this time using deuterated methyl methacrylate monomer (rather than fluorescein) to enable their visualisation on and within the dermis by SRS. The resulting images ( Fig. 7, panels (d–e)), due to the chemical specificity of the Raman CD2 signal, are clear-cut. The blue signal originates exclusively from the deuterated methacrylate monomer incorporated into the nanoparticles confirming that they are exclusively confined to crevasses in the dermis surface and to within the confines of the ablation pore.

  1.    Quantifying Drug Penetration Rates

While the initial effort of the above study was to demonstrate signal detection and spectral separation of pharmaceuticals and excipients within the skin, we have enough information to attempt quantification of ibuprofen, ketoprofen and propylene glycol flux within the pig skin model.  In order to do this, the compound concentration Cx,t at a depth x into the skin at time t, must be first calculated, and Eqtn. W,


,    …W

where M is solute mass released per unit cross-sectional area, and  is the drug’s diffusion coefficient in the skin, first mentioned in Chapter 2, solved for D.  Extracting values for the concentration at two different depths, x1 & x2, for a given time, t, allows us to express:


,    …X

It is then the case that Eqtn. Y,

,       …Y

where K is the skin – solute partition coefficient of the drug, CV is the initial drug concentration in the solute, and G is the total thickness of the stratum corneum, can be used to extract a value for K.

6.4.1. Optical Clearing

One incidental and somewhat helpful effect of the dosing excipient is to increase both the volume and refractive indices of the extracellular spaces within the sample, thereby bringing them into closer alignment with the indices of the cellular scatterers.  This in turn leads to significant enhancement of the imaging depth, as light penetrates deeper into the tissue.  Though this may be considered a happy side-effect of the need to utilise an inert solvent for drug delivery, deliberate use of various substances, specifically for the purpose of improving tissue imaging, has occurred for several years [Insert ref. Zhu et. al. 2013 review paper].  These are known as ‘optical clearing’ agents.

The major tissue optical clearing (TOC) mechanism is considered to be diffusion of the higher-refractive index optical clearing agent (OCA) into intercellular spaces, thereby reducing mismatch between the indices of extracellular fluids and cell components, and thus decreasing light scattering incidents [Insert the three refs. Here].  However, additional mechanisms, including dissociation of collagen fibres [Refs. 1, 2] and temporary replacement of dermal water with OCA [Ref. 3 – 7] have also been suggested.

6.4.2. Depth Correction

Given the variability between individual skin samples, it would seem sensible to obtain depth-correction data for each piece of skin, concurrent with drug/excipient intensity measurements. This can be easily done by switching from the drug to the skin Raman resonance frequency/wavelength (or vice versa) at the designated measurement timepoints, then performing an XYZ stack with otherwise identical measurement parameters.

Initial ‘obvious’ attempts to identify a ‘blanket’ value for signal attenuation with depth coefficient, g, from Eqtn. Z (chapt. 2),

,     …Z

(where S is detected signal intensity, S0 is generated signal intensity in the focal volume, f represents the detection efficiency [constant across data sets for a given wavelength so can be ignored], and L is depth into the skin) using data from undosed pig skin met with problems when applied to dosed skin data, as it tended to overestimate the drug signal with depth, leading to implausibly high values [Appendix X] (up to three times the reservoir concentration) at the deepest points measured.  The main reason for this is believed to be failure to account for the physical effect on skin of the solvent, PG, in which the drugs are dissolved.  PG, of course, has itself sometimes been used both as an excipient for other skin clearing chemicals, and as an optical clearing agent alone [Insert ref.].



Figure X: a) Ketoprofen b) Deuterated propylene glycol.

Fig. X(a) illustrates a saturation of the sample occurring within sixty minutes of dosing.  Concentrations derived from measurements taken at minutes sixty, ninety, and one hundred and twenty all show equilibrium, at least down to a depth of thirty-five microns beneath the skin surface (the maximum depth examined in this timecourse).  The concentrations at minute thirty, however, have not yet reached this equilibrium state, and so can be used to evaluate Eqtn.’s X for D and K, respectively.

Doing so yields values of,

D = 0.2504 µm2s-1,


K = 4.1613

For the undeuterated ketoprofen used in our experiments.

Example calculations can be found in Appendix Z.

Figure Y: Deuterated ibuprofen

6.5.  Discussion

While images of mouse ear dermis obtained with SRS reveal greater structural detail than those from pig dermis of comparable thickness, the acquisition of Raman signal in the forward direction from thinner samples enabled good quality images to be captured. The uptake of formulation constituents by the dermis can cause some swelling of the tissue, but any three-dimensional movement can be tracked by monitoring the CH2 stretching signal from dermis lipids; that is, because Raman scattering permits physiological structures in the dermis to be visualised and used as ‘landmarks’ to decouple chemical diffusion across the dermis from tissue movement.

Although deuteration of chemicals of interest is not an absolute requirement, it can provide significant benefit in the event that there is not a set of wavelengths at which contrast for the dermis, drug and vehicle can be exclusively obtained. In addition, the CD2 wavelength gives rise to extremely low levels of SRS signal from dermis itself, and affords optimum contrast as a result.  The new correction derived in 2.1.2. has allowed values to be extracted which can be used with data-fitting models to derive the skin-solute partition coefficient of a given pharmaceutical and excipient, and the diffusion coefficient of the drug.

SRS offers valuable mechanistic information, such as the transport pathway taken by the penetrant, and the ‘metamorphosis’ of the formulation, including crystallisation of the drug; in other words, key visual insight which is not accessible using alternative techniques. In addition, SRS is an excellent tool with which to investigate the effects of dermis ablation devices, and to determine the disposition of nanoparticles in such porated dermis, in particular those too small to be individually visualised.


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