Vibrational microscopy and imaging of skin: from single cells to intact tissue

Vibrational microscopy and imaging offer several advantages for a variety of dermatological applications, ranging from studies of isolated single cells (corneocytes) to characterization of endogenous components in intact tissue. Two applications are described to illustrate the power of these techniques for skin research. First, the feasibility of tracking structural alterations in the components of individual corneocytes is demonstrated. Two solvents, DMSO and chloroform/methanol, commonly used in dermatological research, are shown to induce large reversible alterations (α-helix to β-sheet) in the secondary structure of keratin in isolated corneocytes. Second, factor analysis of image planes acquired with confocal Raman microscopy to a depth of 70 μm in intact pigskin, demonstrates the delineation of specific skin regions. Two particular components that are difficult to identify by other means were observed in the epidermis. One small region was formed from a conformationally ordered lipid phase containing cholesterol. In addition, the presence of nucleated cells in the tissue (most likely keratinocytes) was revealed by the spectral signatures of the phosphodiester and cytosine moieties of cellular DNA.
Introduction
Technological developments in infrared and Raman microscopic imaging have permitted several new types of biomedical applications. As demonstrated elsewhere in this volume, the diagnosis of pathological states in a wide range of tissues is now feasible, and is being pursued vigorously in laboratories around the world. The vast volume of spectral data generated from imaging experiments permits sophisticated multivariate statistical analysis of data and enables the experimentalist to discern tissue regions where spectral signatures from pathological states differ from those of healthy tissue.
An important attribute of vibrational spectroscopy is the availability of spectra–structure correlations from many tissue components. This extensive background information can provide a useful supplement for biomedical diagnostics. By way of example, Boskey and her collaborators have characterized a new form of brittle bone disease (osteoporosis) based on structural changes in the collagen component of mineralized tissue [1]. This pathological state differs from the more widely known forms of osteoporosis, which are characterized by altered hydroxyapatite levels/structure. The origin of the spectral changes in the collagen component of pathological tissue biopsies was traced to increased levels of collagen cross links and alterations in their spatial distribution. Evidently, the availability of such information facilitates the development of strategies for tracing the origin (i.e., genetic predisposition, external factors, etc.) of the pathology.
Skin is another tissue where the availability of spectra–structure correlations as well as the ability to detect spectral signals from exogenous constituents opens up new possibilities for vibrational microscopic imaging experiments relevant to pharmacology, physiology, and biochemistry [2–4]. In this article, we provide two examples depicting the interplay of vibrational microscopic imaging with spectra/structure correlations.
The first example uses both IR and Raman microscopy to evaluate structural changes in the keratin of single cells (corneocytes) induced by treatment with dimethyl sulfoxide (DMSO) or chloroform/methanol (C/M). These solvents have been widely used in dermatology for studies of permeation enhancement and for extracting lipid from the stratum corneum (SC), respectively. The characterization of DMSO-induced structural changes in keratin will aid in the determination of DMSO permeation pathways and in the evaluation of SC barrier recovery. Assessing C/M’s effect on keratin in isolated corneocytes including the reversibility of potential structural changes, will facilitate the delineation of solvent treatment effects from that of lipid removal.
In the second example, we demonstrate that confocal Raman microscopy along with multivariate (factor) analysis provides the detection of particular skin regions (stratum corneum, epidermis, dermis) as well as endogenous sub-areas (keratinocytes, lipid inclusions) in the epidermis. The availability of spectra–structure correlations permits molecular structure information to be deduced from either the spectra or from the factor loadings.
Materials and methods
Preparation of corneocytes
The Rutgers University Internal Review Board has approved all protocols. Corneocytes were collected by tape stripping (Sellotape, 3M Scotchguard) human forearm skin. The forearm was flushed with water for several minutes prior to tape stripping. The first two strips were discarded and corneocytes from the third tape strip were isolated as follows. Corneocytes were flushed from the tape with HPLC-grade hexane. The hexane/corneocyte suspension was sonicated for approximately 5 min to break up desmosomes and lipid cohesion between individual corneocytes. Corneocytes were then isolated using nitrocellulose membrane filters with a pore size of 0.22 μm (GE Osmonics Labstore, Minnetonka, MN) and resuspended with fresh hexane. This procedure was repeated three times. A small aliquot of this suspension was separately deposited on either CaF2 windows for transmission IR or gold-coated silicon substrates for Raman measurements. Samples were dried overnight under house vacuum prior to spectroscopic measurements. The remaining corneocytes were dried under N2 gas prior to treatment with DMSO or C/M.
Corneocytes were placed on substrates that were directly dipped in pure DMSO for about 1 min. Separate aliquots of harvested corneocytes were transferred to glass vials using 5 mL C/M (2:1 v/v). Non-covalently bound lipids were extracted for 68 h while stirring at room temperature in sealed vials. Corneocytes, collected by centrifugation, were further washed 3 times using fresh C/M before the final suspension was placed on the appropriate substrate. Samples were dried overnight under vacuum prior to spectroscopic measurements. No vibrational bands from residual solvent were observed in corneocyte spectra. Subsequently, corneocytes from both DMSO and C/M treatment were rehydrated on the solid substrates by exposure to 100% relative humidity overnight before a final set of spectra were acquired.
Intact porcine skin preparation
Skin biopsies from Yucatan white, hairless pigs were purchased from Sinclair Research Center, Inc (Columbia, MO). The stratum corneum of the biopsies was washed with water and samples were cut to a size of approximately 1 × 1 cm2, 0.5-cm-thick sections. Sections were placed, SC side up, into a milled brass cell, covered, and sealed with a microscope coverslip for confocal Raman measurements.
IR microspectroscopy of corneocytes
IR spectra were acquired with the PerkinElmer Spotlight system described elsewhere in detail [4]. The instrument has imaging capabilities, but in this study the point mode was used. Since the typical diameter of corneocytes is about 40–60 μm, an aperture size of 40 × 40 μm2 was chosen to optimize the spectral quality. A total of 256 scans was acquired for each spectrum using 4 cm−1 spectral resolution. One level of zero-filling yielded data encoded every 2 cm−1. Approximately 50 corneocytes were examined from each sample preparation.
Raman microspectroscopy
Raman spectra were acquired with a Kaiser Optical Systems Raman Microprobe. The instrument has been described in detail elsewhere [5, 6]. A 785-nm diode laser generates approximately 7–12 mW of single mode laser power at the sample with a spot size of about 2 μm using a ×100 objective. The backscattered radiation illuminates a near-IR CCD (ANDOR Technology, Model DU 401-BR-DD). Spectral coverage is from 100 to 3,450 cm−1 at spectral resolution of 4 cm−1. Following linearization, data are encoded every 0.3 cm−1. Spectra were acquired from 10–15 corneocytes from each sample preparation using an 80-s exposure time, 4 accumulations, and cosmic ray correction. Optical images of corneocytes were acquired with a Leica microscope (Model DMLP) coupled to the Kaiser system with a ×100 objective.
Confocal Raman spectra of intact pigskin were acquired using a 60-s exposure time, 2 accumulations, and cosmic ray correction. A ×100 oil immersion objective was used for the intact pigskin samples. To evaluate axial characteristics of the optical set-up used in these experiments, the following observations were made with the oil immersion lens using protocols previously published [5]. The test sample consisted of five layers of a polymer laminate which incorporated two chemically identical layers (out of the five), each with a thickness of 7.5 μm separated by an intermediate layer of thickness 15 μm. This provides a center-to-center distance of 22.5 μm between the identical layers. Raman intensity profiles show a baseline-resolved width for each of the chemically identical layers of approximately 12–13 μm, a half-width of the intensity profile for each peak of about 8 μm, and a peak-to-peak distance of about 25 μm, in good agreement with the known test sample physical characteristics. The actual axial spatial resolution in the skin samples cannot be directly ascertained.
Data analysis
Grams/32 AI software version 6.0 (Thermo Galactic, Salem, NH) was used for processing individual Raman and IR spectra. Raman spectra were Fourier smoothed (80–90%) and 2-point baselines were applied to spectral regions of interest. This degree of Fourier smoothing could be applied without inducing spectral lineshape distortions. Confocal Raman spectra were analyzed (e.g., factor analysis) and Raman image planes were generated using ISys software version 3.1 (Spectral Dimensions, Inc. Olney, MD). Factor analysis was performed using the ISys score segregation routine. The analysis seeks to detect simple patterns in the relationships between observed variables in order to reduce the dimensionality of the data. Score segregation begins by normalizing PCA scores which may then be sharpened by raising them to a power specified by the acceleration parameter. The default value of 10 was used. Factor loadings are then calculated according to (S′S)−1*S′*X, unless S′S is not invertible, in which case iteration stops, and a factor is deleted before continuing.
Results and discussion
Solvent-induced conformational changes in the keratin of single cells
Although its mechanism of action is not well understood, DMSO has often been utilized as an efficient permeation enhancer for a wide variety of drugs, including antiviral agents, steroids, and antibiotics [7]. It has been suggested to alter protein conformation in the stratum corneum (SC), the outermost layer of skin and the main barrier to permeability. Anigbogu et al. [8] monitored the protein (keratin) α-helix to β-sheet transition in hydrated SC. In addition, Caspers et al. [3] monitored protein structure following DMSO penetration into human SC in vivo by confocal Raman spectroscopy. A broadening of the Amide I contour was noted, although major new spectral features were not resolved. It is noted that DMSO concentrations differed in the two experiments.
IR (1,200–3,600 cm−1) and Raman (800–1,720 cm−1) spectra, shown in Fig. 1a and b, respectively, demonstrate the quality of data obtained from untreated human corneocytes. Several strong bands characteristic of the protein and lipid constituents are observed in each spectrum. The Amide I and Amide II bands observed at approximately 1,650 and 1,550 cm−1 in the IR spectra, respectively, are of particular interest. The Amide I band is also one of the strongest bands in the Raman spectrum. In both types of spectra, the measured Amide I frequency reflects the predominantly helical secondary structure found in epidermal keratin. Absorbance in the Amide I region due to ceramides and other components, some containing amide groups and others C=C, is expected to be low compared to keratin absorbance in corneocytes isolated from the topmost layers of the stratum corneum [6]. Raman spectral quality is very good considering the thickness of an individual corneocyte (0.5–0.8 μm) and is comparable to spectra obtained from much thicker stratum corneum samples and intact skin [2, 5, 8]. It is likely that the observed signal has been enhanced due to a surface-enhanced Raman scattering (SERS) effect between the corneocytes and gold-coated substrate. Raman band assignments pertinent to stratum corneum samples have been listed previously [5, 9, 10].
Fig. 1Vibrational microscopy of human corneocytes. a IR point mode spectrum (1,150–3,600 cm−1 region) acquired using a 40 μm2 aperture of a single corneocyte isolated from the third sequential tape strip applied to human forearm skin. The inset shows the optical image of the corneocyte (bar = 10 μm). b Raman spectrum (800–1,720 cm−1 region) of a similarly isolated corneocyte displaying bands characteristic of the lipid and protein (keratin) components
IR and Raman spectra of isolated corneocytes prior to and following immersion in DMSO, and after an overnight rehydration period (top to bottom trace) are displayed in Fig. 2a and b, respectively. A similar series of IR and Raman spectra acquired from corneocytes treated with C/M are shown in Fig. 2c and d. Both solvents induce several alterations in the spectra. The most significant change in the IR spectra is the appearance of an intense low frequency Amide I component observed for the DMSO-treated sample at 1,626 cm−1 in conjunction with a weak high frequency shoulder at approximately 1,695 cm−1. This pattern is diagnostic for formation of antiparallel β-sheet structure [11]. A redistribution of the maximum intensity in the IR Amide II band from 1,550 to 1,517 cm−1 for the same two spectra is also consistent with β-sheet formation [11].
Fig. 2Solvent-induced conformational changes in the keratin of single corneocytes. a and b IR and Raman spectra, respectively, of an isolated corneocyte prior to and following immersion in DMSO, and after an overnight rehydration period (top to bottom, respectively). c and d IR and Raman spectra, respectively, of a corneocyte prior to and following C/M treatment, and after an overnight rehydration period (top to bottom, respectively). In the IR spectra (a and c), α-helical and β-sheet components of the Amide I (ca. 1,650 cm−1) and II (ca. 1,500–1,550 cm−1) bands are noted. In the Raman spectra (b and d), α-helical and β-sheet components of the Amide I are marked along with disordered constituents at 1,685 cm−1. One component of the Amide III band is marked at 1,240 cm−1
Helix:sheet transitions are well documented for different types of keratin, and have been detected by IR spectroscopy. In particular, Oertel [12] used IR to observe similar DMSO-induced changes in human SC. The current work extends observation of this conformational transition to isolated human corneocytes. It is also noted that, based on the IR Amide I and II band contours, a fair amount of helical structure remains subsequent to DMSO treatment.
The Raman spectrum (Fig. 2b, middle trace) corroborates the β-sheet assignment with the appearance of a high frequency Amide I component at about 1,669 cm−1 and a strong Amide III component at approximately 1,240 cm−1. Both features are within frequency ranges assigned to antiparallel β-sheets [13, 14]. The observed changes in the Amide I region of the Raman spectrum are consistent with a previous report detailing the effects of a series of aqueous DMSO solutions on human SC [8]. Comparisons between individual cells and the SC in the Amide III region are difficult due to overlapped bands from lipid components in the SC sample. In the current work, a second slightly weaker band is also observed for the DMSO-treated cells at approximately 1,270 cm−1. Bands in this region are usually assigned to α-helical structure [13, 14]. Generally, caution is required in associating characteristic frequency ranges with conformation for the Amide III mode given that the NH in-plane bend internal coordinate contributes to a number of modes in the 1,200–1,400 cm−1 region [15] and secondary-structure-dependent coupling occurs between the Amide III and the Cα–H bending vibration [16, 17]. This complexity, in addition to the inherent heterogeneity of corneocytes and variable lipid content of the samples examined herein, precludes an in-depth analysis of the Amide III contour in the current context.
The majority of the DMSO-induced secondary structure change is reversed upon rehydration of the corneocytes as demonstrated by comparison of the top and bottom spectra shown in Fig. 2a and b. Some residual broadening remains on the low frequency side of the Amide I band in the IR spectrum. The average full width at half height for the Amide I band in the IR spectra increased from 44 cm−1 for untreated corneocytes to 52 cm−1 following DMSO treatment and rehydration, indicating that some residual sheet structure may be present. An accurate measure of the Amide I bandwidth in the Raman spectra is precluded by the presence of a weak side chain vibration at approximately 1,608 cm−1. The general reversibility of the conformational change upon hydration was also observed by Oertel [12] for isolated SC.
C/M solutions are commonly used to extract lipids from isolated SC and intact skin samples, but solvent effects on the structure of the remaining constituents have not been detailed. The utility of vibrational spectroscopy for evaluation of keratin secondary structure after lipid removal is somewhat hampered by the native ceramide contribution to the Amide I and II band prior to solvent exposure. In SC samples it is therefore difficult to accurately access the contribution ceramides make to Amide I and II band intensities and position. In the current work, this concern is mostly overcome by acquiring spectra of isolated corneocytes in which the ceramide contribution to the Amide modes is substantially reduced.
Several changes are apparent in the IR and Raman spectra of corneocytes upon exposure to C/M (Fig. 2c and d, compare top to middle traces). Some of the solvent-induced alterations are similar to those just discussed for the DMSO-treated cells, e.g., the low (high) frequency Amide I component in the IR (Raman) spectrum at 1,626 cm−1 (1,670 cm−1) and the appearance of an Amide III band at 1,240 cm−1 in the Raman spectrum. These features provide strong evidence that C/M also induces antiparallel β-sheet formation. Differences are also evident when spectra from the two treatments are compared. Broadening on the high frequency side of the Amide I band in the Raman spectra is much more intense after C/M treatment vs. DMSO (compare the middle trace in Fig. 2d to b). Amide I Raman intensity in this region (ca. 1,685 cm−1) has been assigned to non-hydrogen-bonded disordered structures [13]. The broadening in the Raman Amide I band is not completely reversible upon rehydration of the sample that had been treated with C/M, even though the intensity of the β-sheet component at about 1,670 cm−1 has diminished (Fig. 2d, bottom trace). In addition, upon rehydration, intensity remains in the Amide III band at approximately 1,240 cm−1, although this band also broadens. Finally, measurements of the Amide I bandwidth in IR spectra upon rehydration of several C/M-treated corneocytes are essentially the same (±2 cm−1) as widths measured for untreated cells. The close proximity of Amide I bands arising from helical and random coil structures makes it difficult to differentiate between these two forms. Considering both the Raman and IR spectral data after rehydration, it seems safe to conclude that the majority of keratin exists in a helical form with some disordered structure remaining.
The extent of the C/M-induced protein structural changes has been observed to vary among corneocytes. IR spectra of some cells with significant lipid bands (i.e., those characterized by relatively intense methylene stretching modes near 2,850 and 2,918 cm−1) still exhibit notable protein conformation changes (data not shown), while some cells from which the lipid has been essentially completely extracted do not show significant changes in protein secondary structure. It thus appears that the extent of lipid extraction is not directly related to C/M-induced protein conformational changes. Similar results have been noted in IR studies of DMSO-induced protein conformation changes in SC samples [12]. Additional experiments under conditions of controlled hydration are needed to further explore possible origins of these conformational changes. One explanation for the α-helix to β-sheet structure change in keratin is that DMSO displaces bound water necessary for maintaining secondary structure [12]. DMSO is known to form an association complex with water stronger than that formed between water molecules alone [8]. It may be that C/M destabilizes keratin’s native secondary structure in a manner similar to DMSO, since the majority of the native structure is restored after both solvent treatments when the corneocytes are exposed to high relative humidity.
The importance of the current experiments lies in the fact, that in studies of supposedly native protein structures in the SC, C/M is often used to remove lipid components. As shown in the current work, this procedure is not innocuous, but can induce irreversible changes in protein structure in corneocytes.
Confocal Raman microscopy: delineation of skin regions from factor analysis
As demonstrated above, Raman microscopy complements the information available from IR. Although Raman scattering is inherently weaker than IR absorbance, the technique offers two major advantages for tissue studies. First, the spatial resolution of the method is inherently better than IR. For an excitation wavelength of 785 nm, Rayleigh’s rule for spatial resolution (ΔZ = 1.22λ/2(NA)) predicts a spatial resolution of approximately 0.5 μm for a wavelength (λ) of 785 nm and a numerical aperture (NA) of the ×100 objective of 0.95. In practice, as has been discussed in Materials and methods and elsewhere, the apparent depth and related measured optical properties are degraded by refractive index variations in the tissue [5, 18, 19]. Second, and equally important, spectra may be acquired in a confocal manner. Thus, physical sectioning of samples is not required to obtain molecular structure information from within the tissue. This evidently opens up the possibility for in vivo examination of skin, especially at the low laser powers (ca. 10 mW) used in these experiments. Puppels’ group has demonstrated the utility of confocal Raman determination of water levels in vivo in skin [20].
The benefit of using both a multivariate approach and existing spectra–structure correlations to characterize the microanatomy of skin is demonstrated in the next few figures. An optical micrograph of a 5-μm-thick pigskin section, shown in Fig. 3a, allows delineation of different skin regions. The outermost layered stratum corneum (SC) region appears to be ca. 15-μm thick with the underlying epidermis (ca. 50-μm thick) and dermis discernible. The physical sampling used in the confocal Raman measurement precludes the acquisition of a meaningful micrograph from the same sample; however to allow for comparison, confocal Raman spectra were acquired from a similar pigskin biopsy using approximately the same dimensions. Spectra were acquired from an untreated piece of pigskin, in steps of 2.5 μm in the z direction and 5 μm in the x (or y) direction. The area sampled was 75×40 μm so that 240 spectra (30 × 8) were acquired during the experiment. These were analyzed with factor analysis as described in Materials and methods and as applied to liposome-treated pigskin in a previous report [21]. In the current work, five significant factors were determined as shown in Fig. 3b, while score images for each factor are depicted in Fig. 3c. Three of the score images and corresponding factor loadings in Fig. 3 may be directly correlated with the known regions of skin as follows: Factor 1 corresponds to the SC, Factor 4 to the viable epidermis, and Factor 5 to the dermis. Thus, delineation of the skin regions based on the factor score images is wholly consistent with the optical image. The two remaining factor score images (Factors 2 and 3 ) highlight particular features within the viable epidermis. Examination of the factor loadings or averaged spectra corresponding to those pixels where the respective scores are high provides an indication of the molecular origin of the components. In general, factor plots tend to offer somewhat higher S/N ratios, since they represent a substantial condensation of data. However, it is important to remember that factors include all sources of variance in the data. For example, baseline distortions may appear as “features” in the factor loadings, which however are not related to vibrational modes in the sample. Thus, it is useful to routinely compare factors with appropriately selected spectra. In Fig. 4 an average spectrum of the stratum corneum is overlaid with the average localized to the small spatial region highlighted in Fig. 3c, labeled Factor 2. The average spectrum labeled “ordered lipid” has strong bands near 1,058, 1,130, 1,296, 2,880, and 2,850 cm−1 (Fig. 4b and c). These bands are readily assigned [9, 22] to lipid C–C stretching vibrations (1,058, 1,130 cm−1) of a conformationally ordered (all-trans) chain, lipid CH2 twist (1,296 cm−1) and lipid CH2 stretch (2,850 and 2,880 cm−1). In addition, the bands observed at 605 and 700 cm−1 (Fig. 4a) are indicative of cholesterol [23]. Thus, the small pocket in Fig. 3c (Factor 2) likely contains an ordered lipid phase. The fact that the same bands are present in the average stratum corneum spectrum (Fig. 4, top), albeit at lower intensity, strengthens the assignment. Highly ordered lipid phases (ceramides, fatty acids, and cholesterol) comprise the inter-corneocyte matrix of the stratum corneum. In addition, infrared spectroscopic evidence exists (D.J. Moore, unpublished results) for ordered lipid regions with relatively low protein content in appendages (i.e., pores and hair follicles) well below the stratum corneum. It is possible that the confocal Raman image planes shown in Fig. 3c cut through such an appendage.
Fig. 3a An optical micrograph of a microtomed 5-μm-thick pigskin section acquired using the PerkinElmer Spotlight (bar = 10 μm). The results of factor analysis in the 800–1,150 cm−1 region conducted on a confocal Raman map acquired from intact pigskin. b The five distinct factor loadings generated by the ISys score segregation algorithm (see Materials and methods) are offset and labeled Factor 1–5. c The spatial distribution of factor scores for each of the loadings as marked. Dark blue indicates the lowest score with green, yellow, orange, and red indicative of progressively higher scores. Factor loadings and score images have been assigned to different micro regions in skin as described in the textFig. 4Averaged Raman spectra from within the stratum corneum (top) and lipid (bottom) regions as noted in Fig. 3b and c as Factor 1 and 2, respectively. a The 590–790 cm−1 region with bands assigned to cholesterol marked at 605 and 700 cm−1. b The 800–1,380 cm−1 region displays several bands (1,058, 1,130, and 1,296 cm−1) characteristic for ordered (all-trans) lipid acyl chains. c The CH stretching region (2,800–3,030 cm−1) with methylene symmetric (2,850 cm−1) and asymmetric (2,880 cm−1) stretching modes noted. The frequencies and relative intensity of the bands are consistent with well-ordered lipid acyl chains
Finally, Factor 3 (Fig. 3b), whose score image reveals several small-localized pockets in the epidermis, contains one broad spectral feature centered at approximately 1,090 cm−1 that is not present in the other factor loadings. Based on both the position of this particular band and the spatial distribution of the factor score, the band is tentatively assigned to the phosphodiester mode of DNA [24, 25]. Since the identification of DNA is of interest, as it would reveal the location of nuclei within keratinocytes or other cells in the tissue, an additional strategy was employed to acquire better spectra from the epidermis.
The experiment, whose results are depicted in Fig. 5, was carried out as follows. The top ≈20 μm of a pigskin section, including most of the SC, were removed by tape stripping prior to confocal Raman imaging. This allows for the acquisition of higher quality spectra in deeper regions of the viable epidermis. The Raman experiment was also carried out with more data points/unit area (2-μm step size) providing a closer examination of the tissue compared to the sample presented in Fig. 3. The presence of cell nuclei was revealed by factor analysis conducted over two wavenumber regions, each separate analysis restricted to compute only two factor loadings. An additional DNA band assigned to cytosine is detected in the factor loadings shown in Fig. 5b, which also strengthens the previously identified phosphodiester DNA band assignment at about 1,090 cm−1 [24, 25]. As expected, in the tape stripped sample, the phosphodiester band is again observed (Fig. 5c). The score image shown in Fig. 5a is derived from the factor with the strong cytosine band and is essentially the same as the score image generated from the phosphodiester factor (not shown). Thus, the image shown in Fig. 5a, depicting keratinocyte or other epidermal cell (e.g., melanocyte) nuclei, demonstrate the ability of confocal Raman to detect DNA in intact skin tissue.
Fig. 5The results of factor analysis conducted in two separate spectral regions for a confocal Raman map of pigskin after stratum corneum removal by tape stripping. Both analyses were performed allowing the generation of only two factors. In each case, one factor contained a spectral feature specific to DNA and the other mapped in a fairly uniform way throughout the epidermal region. a Factor score image for the factor-labeled nuclei shown in (b) with red corresponding to the highest score and dark blue to the lowest. b Factor loadings for the 600–820 cm−1 region with a distinctive vibrational band at approximately 785 cm−1 assigned to cytosine. c Factor loadings for the 800–1,150 −1 region with a second characteristic DNA vibrational band due to the phosphodiester backbone stretching mode, at about 1,090 cm−1 noted
Conclusions
The current experiments highlight some unique advantages of vibrational microscopic imaging. An understanding of the spectroscopy of the various tissue components permits insight into the molecular origins of the images generated from factor analysis. Thus, identification of the lipid pocket in Fig. 3c (Factor 2) and of the cell nuclei (as detected from the DNA spectral signatures) in Fig. 3c (Factor 3) and Fig. 5a were greatly facilitated by the availability of Raman spectra of lipids and of DNA, respectively. In addition, the availability of correlations between lipid spectra and chain conformation, permitted assignment of the lipid inclusions as having arisen from ordered lipid phases. The biological relevance of this observation remains to be determined. From a pharmacological perspective, the detection of cell nuclei (most likely keratinocytes) within intact skin permits co-localization experiments to begin to be designed. For example, for classes of drugs targeted to these cells, confocal Raman measurements will permit determination of whether the putative therapeutic agent reaches its intended target.
In addition, although all microscopic methods can obviously readily detect single cells, the ability of IR- or Raman-based methods to detect conformational changes within a particular component of a single isolated cell is unique. The example we have chosen in Fig. 2 is of practical importance in skin research, since solvents such as DMSO or chloroform/methanol are used for permeation enhancement and lipid extraction, respectively. In each case these solvents were found to be far from innocuous. Each induced large conformational changes in the cellular keratin. This fact had been known previously from spectra of intact SC, but two aspects of the current measurements are novel. The observation of IR data from single cells provides a sharper spectral assignment of the bands. We note for example that ceramides, a major component of the SC, have strong bands arising from the amide bond located in these lipids. These modes contribute significantly to the spectra of the intact SC and overlap protein Amide I and II vibrations; however, they are much reduced in relative intensity in spectra of cells. Thus, studies of the reversibility of the solvent-induced conformational changes benefit from the reduced intensity of ceramide interference.
Finally, it is feasible to begin to examine biochemical heterogeneity at a single cellular level. Issues such as the relationship of protein structure (either native or solvent-modified) to hydration levels or to the levels of natural moisturizing factors, in single cells may be profitably probed. In addition, changes in the spectra of cells from pathological states may be examined.

corneocytes
confocal raman
epidermis
protein secondary structure
infrared imaging


RGD-avidin–biotin pretargeting to αvβ3 integrin enhances the proapoptotic activity of TNFα related apoptosis inducing ligand (TRAIL)

Recombinant TNF-related apoptosis-inducing ligand (TRAIL) is considered a powerful and selective inducer of tumor cell death. We hypothesize that TRAIL’s potential as anticancer agent can be enhanced further by promoting its accumulation in tumor tissue. For this purpose, we developed TRAIL complexes that bind to angiogenic endothelial cells. We employed an avidin–biotin pretargeting approach, in which biotinylated TRAIL interacted with RGD-equipped avidin. The assembled complexes killed tumor cells (Jurkat T cells) via apoptosis induction. Furthermore, we demonstrated that the association of the RGD-avidin-TRAIL complex onto endothelial cells enhanced the tumor cell killing activity. Endothelial cells were not killed by TRAIL nor its derived complexes. Our approach can facilitate the enrichment of TRAIL onto angiogenic blood vessels, which may enhance intratumoral accumulation. Furthermore, it offers a versatile technology for the complexation of targeting ligands with therapeutic recombinant proteins and by this a novel way to enhance their specificity and activity.
Introduction
TNF ligands have strong potential as anti-tumor agents, acting in different ways on either tumor cells or tumor vasculature, or by promoting the induction of tumor-directed immune responses [1]. The broad clinical use of TNF ligands is prohibited however due to lack of tumor selectivity, causing various side effects. A recently identified TNF family member is TNF related apoptosis-inducing ligand (TRAIL) [2]. Many tumor types are sensitive to TRAIL or become sensitive after treatment with chemotherapeutic drugs [3]. Soluble TRAIL variants showed no cytotoxic effects on most primary human cells and were well tolerated in mice and chimpanzees [4, 5]. However, the selectivity of TRAIL for tumor cells over normal cells is lost when TRAIL forms multimeric complexes. For example, it has been observed that cross-linked TRAIL induces apoptosis more efficiently than non-aggregated trimeric TRAIL [6], but such cross-linked TRAIL also caused cell death in normal hepatic cells [7]. It seems therefore attractive to design clustered TRAIL variants with enhanced specificity for tumor tissue. The increased tissue retention of cellular targeted TRAIL variants may counterbalance or prevent side effects elsewhere in the body. In the present paper, we investigated the development of TRAIL complexes equipped with targeting ligands directed to the tumor vasculature. Enhanced retention of TRAIL complexes at the tumor vasculature will enhance their accumulation in the tumor tissue, thereby favoring the desired therapeutic activity and reducing potential toxicity.
Several recent papers described the development of recombinant TNF family members with enhanced target specificity. One approach is to modify the intrinsic specificity of TRAIL for its target Death-receptor 5 specific by substituting amino-acids located in the receptor binding interface of TRAIL [8]. Development of chimaeras is another successful approach. Linkage of TRAIL to scFv antibody fragments yielded constructs which bind to additional surface-exposed receptors on tumor cell types [9–11]. Similarly, other TNF family members like Fas-ligand and TNFα (have been conjugated to tumor-cell directed targeting ligands, yielding chimeric proteins with full or even enhanced therapeutic activity [11–15].
Although the recombinant approach has been applied successfully for the production of chimeric proteins, low production yields may hamper the feasibility of such a strategy. Especially when multiple targeting ligands are introduced into a protein subunit, which is preferred for high avidity binding, aberrant folding of chimeric proteins can occur. In the current study we therefore pursued a chemical approach, in which TRAIL was biotinylated and complexed to avidin equipped with multiple targeting ligands. Avidin–biotin complexation is a widely accepted approach in diagnostic applications that employ immunodetection steps, and is also considered an attractive strategy for the preparation of tumor targeted complexes, especially in so-called pretargeting approaches in which the complexes are assembled in vivo at the cell surface of tumor cells. Commonly, such strategies employ biotinylated tumor-specific antibodies and a therapeutic or diagnostic cargo coupled to avidin [16, 17].
The approach presented in this paper differs to the above-discussed strategies in several aspects. First, we now equipped TRAIL with targeting ligands directed to angiogenic endothelium rather than to tumor cells, allowing an enhanced binding to the tumor blood vessels. The vascular wall is in direct contact with the systemic circulation, which makes it an easily attainable target for therapeutic proteins. Furthermore, intratumoral pressure, which hampers the penetration of macromolecules into tumor tissue will not affect the homing to of RGD-equipped complexes to the angiogenic endothelium.
Second, we have prepared a different type of targeted avidin that can interact directly via covalently attached RGD-groups with tumor endothelial cells. Thus, the developed avidin–biotin-TRAIL complexes will home to angiogenic endothelium without the need of complexation with a biotinylated antibody, thereby leaving its biotin-binding sites available for complexation with the biotinylated TRAIL. The functionalization of avidin with RGD-homing ligands was carried out using a cyclic RGD-peptide with high affinity and specificity for αvβ3 integrin on angiogenic endothelium [18]. We furthermore applied a polyethylenglycol (PEG) linker for coupling of the RGD-ligand, which brings several advantages, such as a reduced non-specific binding of the avidin and reduced immunogenicity [19]. The now developed RGDPEG-avidin conjugates contain the RGD homing ligand attached to the distal end of the PEG groups.
In order to minimize loss of functional TRAIL activity, we have compared two different biotinylation strategies that react at different functional groups in the protein. We used a conventional Biotin-NHS reagent that binds to primary amino groups in the protein, and we employed the Biotin-ULS (ULS: universal linkage system) reagent, which reacts at methionine residues. Figure 1 shows a schematic representation of the developed complexes and their interaction with target cells. The products were evaluated for their association with αvβ3 integrin expressing endothelial cells and for their antitumor activity in both endothelium and cultured tumor cells. Our results demonstrate that recombinant TRAIL can be complexed with endothelial specific targeting ligands in such a way that it maintains its extensive cell killing activity either in solution or when displayed on the surface of endothelial cells.
Fig. 1Schematic representation of RGD-avidin:TRAIL complexes. Soluble recombinant human TRAIL was modified with biotin groups via two different reagents that either react at lysine or methionine residues. Biotinylated TRAIL was complexed to an avidin-based targeting device equipped with approximately 5 RGD-peptide ligands per avidin molecule. A hydrophilic PEG-linker was used to append the RGD-peptides to avidin. The assembled complexes can bind to tumor blood vessels via RGD//αvβ3-integrin interactions, while apoptosis can be induced via TRAIL/DR ligation
Materials and methods
Preparation and characterization of RGDPEG-avidin
Avidin was modified with heterobifunctional PEG groups that can react with primary amino groups in avidin, after which RGD-peptide groups can be conjugated to their distal ends. The hydrophilic PEG groups of 3.4 kDa will furthermore reduce non-specific protein interactions.
Briefly, avidin (2 mg, 32 nmol, Sigma) was dissolved in PBS at 10 mg/ml and treated with a 50-fold molar excess of the heterobifunctional vinylsulfone-polyethylenglycol-N-hydroxysuccinimide ester (1 mg, 292 nmol, VNS-PEG-NHS; Nektar, Alabama, USA), which had been dissolved in water at a concentration of  20 mg/ml. The PEG linker was added drop wise. The reaction mixture was protected from light with tin foil and incubated for 1 h at room temperature on a spiramix rollerbank. Meanwhile, the RGD peptide c(RGDf(ε-S-acetylthioacetyl)K) (Ansynth Service, Roosendaal, The Netherlands) was dissolved at 10 mg/ml in a 1:4 acetonitrile/water mixture. The peptide was added drop wise to the reaction mixture at a molar ratio of 55:1, after which freshly prepared hydroxylamine was added to a final concentration of 50 mM. The reaction was carried out overnight at room temperature while protected from light. Remaining VNS groups were quenched by addition of cysteine (55:1 molar excess over the amount of avidin), after which the product was purified by size exclusion chromatography on a Highload superdex 200 column (Amersham Biosciences), pre-equilibrated in PBS. The final product RGDPEG-avidin was stored in PBS at −20°C.
RGDPEG-avidin conjugates were characterized for protein content by BCA protein assay (Pierce). Coupling of RGDPEG groups to avidin was demonstrated by SDS-PAGE performed on a mini-Protean II system (Bio-Rad) using 12% Ready Gels (Bio-Rad, The Netherlands). Duplicate gels were either stained for protein (Coomassie Brilliant Blue staining) or blotted on a PVDF membrane (Roche, Mannheim, Germany). The membrane was blocked with BSA and subsequently incubated with an in-house raised anti-RGD polyclonal rabbit antiserum specific for the conjugated RGD-peptide [20]. Blots were further developed with goat-anti-rabbit peroxidase (GARPO)/DAB detection. The grade of RGD-pegylation was furthermore determined by MALDI-TOF analysis as described previously [21].
Expression and purification of recombinant human His-tagged TRAIL
A cDNA corresponding to human soluble TRAIL (amino acids 114–281) was cloned downstream of the hexahistidine (His) tag sequence in pET15B (Novagen) using NdeI and BamHI restriction sites. Escherichia coli BL21 (DE3) (Invitrogen) was subsequently transformed with the His-TRAIL construct. The transformed host was grown in shake flasks at four liter scale as described previously [22]. The culture was centrifuged at 2,500 rpm and the supernatant was discarded. The isolated pellet was resuspended in three volumes of extraction buffer (20 mM phosphate buffer pH 8 containing 200 mM NaCl, 10% (v/v) glycerol and 7 mM β-mercaptoethanol).
Cells were disrupted using sonication and extracts were clarified by centrifugation at 40,000 g. Subsequently, the supernatant was loaded on a cobalt-charged IMAC TALON column (Clontech) and His-TRAIL was eluted with a step gradient of extraction buffer containing 0.5 M imidazole. Trimeric His-TRAIL was purified by size-exclusion chromatography on a Hiload Superdex 75 column (Amersham Biosciences), to remove impurities. The final product was flash frozen and stored at −80°C in storage buffer (phosphate buffer pH 7.4 containing 150 mM NaCl, 3.5 mM DTT and 20 μM ZnSO4). Purified protein was >95% pure as determined by colloidal Coomassie Brilliant Blue-stained SDS–PAGE gel.
Preparation and characterization of bio-NHS-TRAIL and bio-ULS-TRAIL
We studied two different linkage strategies to introduce biotin into His-TRAIL, by applying either a conventional biotinylation reagent that reacts with primary amino groups in the protein (biotin-NHS), or one that reacts to methionine residues (biotin-ULS™, ULS = universal linkage system). Such an alternative strategy may prove less harmful to the therapeutic activity of TRAIL, depending on the extent to which side chain residues in the active domains are modified. Biotin-ULS was provided by Kreatech Biotechnology (Amsterdam, The Netherlands). All syntheses were performed in a 10:1 molar ratio of biotinylation reagent over TRAIL.
His-TRAIL in storage buffer was extensively dialyzed against PBS containing 10% glycerol at 4°C using Slide-A-Lyzer dialysis cassettes (10,000 MWCO, Pierce) to remove DTT and other components of the storage buffer that can interfere in the reactions. Typically, the purified His-TRAIL (500 ug, 7.6 nmol) at 0.7 mg/ml was mixed either with Sulfo-NHS biotin (Pierce) (10 mg/ml in DMF) or with biotin-ULS (10 mg/ml in 20 mM NaCl), after which the mixtures were incubated for 4 h at 37°C. Unreacted biotinylation reagent was removed by dialysis against PBS at 4°C. The products were sterilized by filtration via disposable 0.2 μm filters and stored at −20°C.
Biotin-TRAIL conjugates were characterized for protein content using BCA protein assay (Pierce). The relative number of coupled biotin molecules was determined by anti-biotin ELISA. Wells were coated with serial dilutions of the biotinylated proteins for 1 h at room temperature, washed with PBS containing 0.05% Tween20 and incubated for 1 h at room temperature with streptavidin-peroxidase complex (Dako, 1:2500 in PBS), followed by a standard incubation with OPD. The protein concentration at which 50% of the maximum absorbance was measured (EC50) was calculated by nonlinear regression (Graphpad Prism), and used to calculate relative biotin:protein ratios in each conjugate. The grade of biotinylation was furthermore assayed by MALDI-TOF analysis as described above for RGDPEG-avidin.
Cells
Jurkat T cells were cultured in RPMI 1640 culture medium (containing 25 mM HEPES and l-glutamine), supplemented with 10% heat-inactivated FCS (Bodinco, Alkmaar, The Netherlands), 1 mM sodium pyruvate, 2 mM l-glutamine, 0.5 mM 2-ME, and 0.1 mg/ml gentamycin sulfate obtained from BioWhittaker, and 0.02 μg/ml fungizone. (Bristol-Meyers, Woerden, The Netherlands)
HUVEC were isolated by the UMCG Endothelial Cell Facility by previously described methods [23]. Primary isolates were cultured in 1% gelatin-coated tissue culture flasks (Corning, Costar, The Netherlands) at 37°C under 5% CO2/95% air. The culture medium, hereafter referred to as EC medium, consisted of RPMI 1640 (BioWhittaker, Verviers, Belgium) supplemented with 20% heat inactivated fetal calf serum (Integro, Zaandam, The Netherlands), 2 mM l-glutamine (Invitrogen, Breda, The Netherlands), 5 U/ml heparin (Leo Pharmaceutical Products, Weesp, The Netherlands), 100 U/ml penicillin (Yamanouchi Pharma, Leiderdorp, The Netherlands, 100 μg/ml streptomycin (Radiumfarma-Fisiopharma, Italy), and 50 μg/ml endothelial cell growth factor supplement extracted from bovine brain. After attaining confluence, cells were detached from the surface by trypsin EDTA (0.5/0.2 mg/ml in PBS) treatment and split in 1:3 ratio. Cells were used up to passage four.
Binding of RGDPEG-avidin to HUVEC
Binding affinity of RGDPEG-avidin to αvβ3-integrin expressed on the surface of HUVEC was determined by competitive displacement of 125I-labeled Echistatin, which is a known ligand for αvβ3 [24]. Confluent monolayers of HUVEC in 24-well plates (Costar) were incubated with 100,000 cpm 125I-Echistatin either in the presence of excess of RGDPEG-avidin, free RGD-peptide or avidin as competitors (competitors all in a concentration of 10 μM), or a serial dilution of RGDPEG-avidin and free RGD-peptide. Competition experiments were carried out with divalent cation containing binding buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM CaCl2, 1 mM MgCl2, 1 mM MnCl2 and 1% BSA) at 4°C for 4 h. Cells were washed three times with binding buffer and lysed with 1 M NaOH. Radioactivity was counted in a Packard RIASTAR multiwell gamma counter (GMI, Minnesota, USA). Data were analyzed by non-linear regression using the GraphPad Prism program.
Complexation of biotinylated TRAIL with RGDPEG-avidin
We studied the complexation of biotinylated TRAIL with RGDPEG-avidin in an ELISA-like experimental setup. Wells were coated with 50 ng of avidin or RGDPEG-avidin overnight at room temperature, blocked with BSA (1% in PBS) and incubated with 10 ng of biotinylated TRAIL for 2 h at room temperature. After the wells were washed with PBS/0.05% Tween20, the amount of TRAIL associated with the avidin was detected by anti-TRAIL immunodetection (anti-TRAIL antibody 2E5, Alexis, Breda, The Netherlands) for 1 h, followed by standard detection with GARPO/OPD.
Apoptotic activity of biotinylated TRAIL
The apoptosis inducing activity of the biotinylated products was evaluated on Jurkat T cells and on endothelial cells, using a cell viability assay and a caspase activity assay. Experiments were conducted with the single cell types, Jurkat tumor cells or HUVEC, or with a combination of tumor and endothelial cells.
Assessment of TRAIL activity by MTS cell viability assay
Briefly, cells were seeded in flat-bottom 96-well plates at a density of 3 × 104cells/well (Jurkat) or 1 × 104cells/well (HUVEC) in 100 μl medium supplemented with the indicated compounds. Concentrations of TRAIL, bio-NHS-TRAIL and Bio-ULS-TRAIL ranging from 1 μg/ml to 1 ng/ml were tested on Jurkat. Only one concentration (100 ng/ml) was tested on HUVEC. The MTS cell viability assay (Promega) was performed after 48 h of incubation, according to the manufacturer’s recommendations. Experiments were performed in triplicate.
Assessment of TRAIL activity by caspase 3/7 assay
We investigated the induction of apoptosis in Jurkat T cells using a caspase 3/7 activity assay (Promega) employing luminescence detection. Different experimental settings were evaluated either reflecting direct incubation of the products with tumor cells, or settings in which biotinylated TRAIL was complexed to RGDPEG-avidin and/or anchored onto endothelial cells.
First, we determined pro-apoptotic effects of non-complexed TRAIL. About 5 × 103 Jurkat cells were incubated with biotinylated TRAIL at a concentration of 100 ng/ml. Experiments were carried in 96-well plates in 50 μl medium supplemented with the indicated compounds for 4 h, after which 50 μl of caspase 3/7 substrate was added to the cells, followed by luminescence detection with a microplate luminometer according to the provider’s instructions.
Next, we determined proapoptotic effects of RGDPEG-avidin:TRAIL complexes in Jurkat cells. These experiments were performed in the presence of endothelial cells, to allow adherence of the RGD-equipped complexes to αvβ3 integrin expressing cell layers. HUVEC were preincubated with 10 μg/ml RGDPEG-avidin for 2 h at room temperature after which the excess of non-cell associated RGDPEG-avidin was removed by washing with PBS. Control experiments included RGDPEG-avidin treated HUVEC that were not washed and cells to which no RGDPEG-avidin was added. All wells were supplemented with biotinylated TRAIL (100 ng/ml in culture medium), after which Jurkat cells (5 × 103 cells in 50 μl of medium) were added. Cells were incubated for 4 h at 37°C, after which caspase activity was assayed as described above. Experiments were conducted in triplicate.
Statistical analysis
Statistical analysis was performed using Student’s two-tailed t-test, assuming equal variances. Difference were considered to be significant when P < 0.05 unless otherwise stated.
Results
Preparation and characterization of RGDPEG-avidin
We modified avidin with PEG-linked RGD-peptides to obtain an avidin molecule which could bind to angiogenic endothelial cells. Successful synthesis of RGDPEG-avidin was proven by several analytical techniques. SDS-PAGE analysis (Fig. 2a) showed a single band for avidin of approximately 15 kDa, corresponding to the weight of one of its four subunits. Different protein bands were observed for RGDPEG-avidin with molecular weights corresponding to the unmodified subunit and to subunits modified with one or multiple RGDPEG moieties. Second, Western blot of the product showed positive staining of these bands by an anti-RGD-peptide antiserum. Due to the size heterogeneity of the PEG polymer, the RGD-band stained in a broad streak rather than a discrete band (Fig. 2b). Of note, the tetrameric band of avidin was not observed due to the denaturating conditions of SDS-PAGE. Third, we analyzed the product by MALDI-TOF mass spectrometry (Fig. 2c), which showed peaks corresponding to avidin subunits modified with RGDPEG to a different degree. From the average molecular weights (Mn) we calculated a relative derivatization ratio of 1.4 RGDPEG groups per avidin subunit, or a total of 5.6 RGDPEG groups per tetrameric avidin.
Fig. 2Characterization of RGDPEG-avidin. Panel A and B: SDS-PAGE separation followed by CBB protein staining (a) or Western blotting and anti-RGD detection  (b). lane 1: MW marker, lane 2: avidin, lane 3: RGDPEG-avidin. Unmodified avidin was separated into its 15 kDa subunits, while RGDPEG-avidin was disassembled into different bands corresponding to subunits with 0, 1, 2 or 3 attached RGDPEG groups. The intrinsic heterogeneity of PEG polymers can be observed in the broadly stained band of the RGDPEG-avidin Western blot. Panel C: MALDI-TOF analysis of RGDPEG-avidin. Upper line: RGDPEG-avidin, bottom line: avidin. During ionization, the tetrameric proteins have disassembled into subunits, yielding a single peak for avidin and multiple peaks for RGDPEG-avidin corresponding to modification degree 0, 1, 2, 3, 4
Next, we investigated the interaction of RGDPEG-avidin with HUVEC, which express αvβ3-integrin when being at a subconfluent state. To determine binding affinity of RGDPEG-avidin for αvβ3-integrin, we studied its capability to compete with the αvβ3-integrin ligand 125I-echistatin. While RGDPEG-avidin completely displaced 125I-echistatin from the cells, similar to the free RGD-peptide, avidin did not affect binding of the radiolabeled peptide (Fig. 3a). The sigmoid displacement curve with RGDPEG-avidin (Fig. 3b) demonstrated a 2.9-fold increase in affinity (IC50 = 134 nM) as compared to the single RGD-peptide (IC50 = 383 nM).
Fig. 3Association of RGDPEG-avidin with endothelial cells. Subconfluent HUVEC were incubated with radiolabeled echistatin, which is a known ligand for αvβ3 integrin, and RGDPEG-avidin, free RGDpeptide or unmodified avidin. Panel A: Binding specificity. Echistatin was displaced by excess of RGDPEG-avidin or free RGD-peptide (both at 10 μM) but not by excess of avidin. *P < 0.05. Panel B: Affinity of binding. Echistatin binding was displaced by different concentrations of the added competing ligands, showing the increased affinity of RGPEG-avidin over the single RGDpeptide ligand
Preparation and characterization of biotinylated TRAIL
Since the modification of side-chain residues may diminish the activity of a therapeutic protein, we studied two different biotinylation strategies that target chemical modifications at different amino acid residues, i.e., at lysine groups (biotin-NHS) or methionine residues (biotin-ULS). Such different labeling positions at the surface of the protein may correlate to differences in pharmacological activity of the biotinylated proteins. Anti-biotin ELISA (Fig. 4a) demonstrated a higher biotin/TRAIL modification of biotin-NHS over biotin-ULS, which is in line with the lower abundance of methionine residues as compared to the number of lysine residues. We tried to confirm these data by analysis of the biotinylation extent by MALDI-TOF mass spectrometry, which provided reliable data on the average number of incorporated RGDPEG units in avidin. Although this approach succeeded for biotin-NHS-TRAIL (Fig. 4b), demonstrating the incorporation of about 1 biotin group per subunit of TRAIL, MALDI-TOF analysis failed to detect biotin-ULS modification of TRAIL. Most likely, the mass ionization step induced the dissociation of biotin-ULS from the protein. From the collective results, we concluded that both strategies had yielded biotinylated TRAIL with approximately 1–3 biotin groups per protein (summarized in Table 1).
Fig. 4Characterization of biotinylated TRAILs. Panel A: Anti-biotin ELISA with streptavidin-HRP. Signals were corrected for background and expressed relative to the maximum detected intensity. Panel B: MALDI-TOF analysis of TRAIL and biotin-NHS-TRAILTable 1Characteristics of biotinylated TRAIL variantsRelative biotin content (Biotin/TRAIL)aActivity in cell viability assay (IC50 in ng/ml)bActivity in caspase 3/7 assayc (%)TRAIL–40100Bio-NHS-TRAIL1.612335Bio-ULS-TRAIL0.860126aExpressed as mole biotin bound per mole of trimeric TRAIL. Relative biotin content was determined by MALDI-TOF analysis (bio-NHS-TRAIL) or ELISA (BIO-ULS-TRAIL)bAs determined after 48 h of incubation with Jurkat cellscAs determined after 4 h of incubation at 10 ng/ml with Jurkat cells
We studied complexation of biotinylated TRAIL with RGDPEG-avidin that had been coated in 96-well plates. Highest binding was observed for Bio-NHS-TRAIL, probably due to its higher biotin content, while bio-ULS-TRAIL showed approximately 40% less binding (Fig. 5). Unmodified His-TRAIL showed non-specific binding to RGDPEG-avidin as well, despite the absence of biotin groups in this product. Non-specific binding of TRAIL to RGDPEG-avidin was however far lower than binding to unmodified avidin, most likely due to the PEGylation (data not shown).
Fig. 5Association of biotinylated TRAIL with RGDPEG-avidin. 96-well plates were coated overnight with 50 ng RGDPEG-avidin and incubated with indicated compounds for 2 h. TRAIL binding was assessed by anti-TRAIL immunodetection and expressed relative to the highest signal obtained with biotin-NHS-TRAIL. Experiments were performed in triplicate
Cytotoxic and proapoptotic activity of TRAIL complexes
To test whether our products were functionally active, i.e., capable of inducing cell death and apoptosis, we assessed their activity on Jurkat T leukemic cells and HUVEC endothelial cells. Figure 6a shows the dose-dependent cytotoxic effects at Jurkat cells after treatment for 48 h. Both His-TRAIL, Bio-NHS-TRAIL and Bio-ULS-TRAIL effectively killed Jurkat cells with IC50 values between 40 ng/ml and 123 ng/ml (Table 1). While the Bio-NHS modification resulted in a considerable reduction in activity compared to the unmodified protein, Bio-ULS-TRAIL displayed almost similar activity as unmodified TRAIL. The pro-apoptotic activity of His-TRAIL and biotinylated TRAIL derivatives in Jurkat cells was furthermore determined by analyzing caspase 3/7 activity, as shown in Fig. 6b. These experiments confirmed the relative higher activity of bio-ULS-TRAIL versus bio-NHS-TRAIL, and the absence of caspase induction by RGDPEG-avidin (data not shown). In contrast to Jurkat cells, no cytotoxic effects were observed when HUVEC were incubated with the products (Fig. 6c).
Fig. 6TRAIL activity assays. Panel A: Cell viability of Jurkat T cells after 48 h of incubation with indicated compounds. Cell viability was assessed by MTS assay. Experiments were performed in triplicate. Panel B: Caspase 3/7 activity in Jurkat T cells after 4 h of incubation with 100 ng/ml of the indicated TRAIL derivatives. RLU: relative light units. Panel C: Cell viability (MTS assay) of HUVEC after 48 h of incubation with 100 ng/ml of indicated compounds
Finally, we investigated the proapoptotic effects of RGDPEG-avidin:TRAIL complexes in Jurkat cells. These experiments were performed in the presence of endothelial cells, to allow adherence of the RGD-equipped complexes to αvβ3 integrin expressing cell layers. First, we determined the effects of avidin:TRAIL complexes formed in the presence of an excess of RGDPEG-avidin. As can be observed in Fig. 7, such complexes  with both types of biotinylated TRAIL showed similar activity to the non-complexed biotinylated TRAIL derivatives (striped bars versus white bars, respectively). Furthermore, in agreement with Fig. 6b, Bio-NHS-TRAIL showed a reduced activity as compared to unmodified TRAIL, while Bio-ULS-TRAIL retained its activity.
Fig. 7Proapoptotic activity of RGDPEG-avidin:TRAIL complexes. Assays were performed in 96-well plates seeded with HUVEC that were subsequently incubated at room temperature with RGDPEG-avidin (10 μg/ml)  and biotinylated TRAIL (100 ng/ml), to allow assembly of the complexes. Next, Jurkat T cells were added followed by 4 h of incubation at 37°C, after which the activation of caspase 3/7 was determined by a luminescence based assay. Experiments were carried out in the absence of RGDPEG-avidin (control experiment, white bars), in the presence of excess of RGDPEG-avidin (striped bars, 100-fold molar excess over biotinylated TRAIL) or in wells incubated with RGDPEG-avidin that had been washed before addition of biotin-TRAIL and Jurkat cells (black bars). *P < 0.05 versus TRAIL; #P < 0.05 versus biotin-TRAIL in the absence of RGDPEG-avidin
A striking result was observed when the excess of RGDPEG-avidin was removed before addition of TRAIL and Jurkat cells, leaving only endothelial-associated RGDPEG-avidin available for complexation with biotinylated TRAIL. This procedure significantly potentiated the proapoptotic effect for both Bio-NHS-TRAIL and Bio-ULS-TRAIL, the latter even beyond the activity of unmodified TRAIL.
Discussion
In the present study we describe the development and functional evaluation of RGD-avidin:TRAIL complexes. These targeted complexes of TRAIL showed enhanced binding to endothelial cells and maintained their pro-apoptotic activity. As a result of the binding to angiogenic endothelial cells of these complexes, an improved tumor accumulation and enhanced residence within tumor tissue can be expected. Such an improvement in the pharmacokinetic profile of TRAIL may further enhance its therapeutic efficacy and enable a reduction in dosing frequency.
αvβ3 integrin is a validated target for drug delivery and imaging approaches directed to angiogenic endothelium. One of the most popular ligands for αvβ3 integrin is the cyclic RGD-peptide cyclo(RGDfK). Due to the special constrained conformation of the cyclic RGD-peptide and the d-phenylalanine amino acid, its binding motif adopts a high affinity conformation for αv-integrins [25]. The cyclo(RGDfK) peptide has been used successfully for the delivery of a wide range of drug or gene carrying systems (e.g., liposomes, polymers, adenoviruses) [18]. Since the cyclopeptide is an unnatural peptide, it can not be encoded by a DNA sequence and requires chemical synthesis and hence chemical coupling techniques to prepare multivalent targeting devices. The present paper describes a versatile approach in which multivalent RGD-avidin is complexed with biotinylated TRAIL. The RGD-peptides are linked to avidin via bifunctional PEG linkers, thereby surrounding the avidin by a hydrophilic PEG shell tethered with targeting ligands. PEGylation is a widely accepted methodology for improvement of therapeutic proteins, and affects both the immunological properties as well as the renal filtration of proteins [26]. The functionalization of avidin with RGD-PEG groups thus will combine several advantageous properties. RGDPEG-avidin displayed nanomolar affinity for αvβ3 integrin expressing endothelial cells, which is in good agreement with the data obtained with other RGD-equipped proteins prepared in our group [27].
We have compared two different biotinylation procedures, which react at different amino acid side chain residues. While biotin-NHS couples at lysine residues, biotin-ULS binds at methionines residues in TRAIL. The methionines in TRAIL are not solvent exposed, as can be derived from the crystal structure of TRAIL [28]. Therefore, the most likely binding site for biotin-ULS is the methionine in the HIS-tag of the recombinant protein. In contrast, most lysine residues are readily accessible, including ones at positions that will interface with the Death receptor. This could explain why Biotin-ULS-TRAIL better maintained its proapoptotic activity than biotin-NHS derivatized TRAIL. Whether the greater loss in activity of biotin-NHS-TRAIL is due to the relative higher extent of derivatization or to modifications of lysine residues in the activity domains of protein can not be concluded from our data.
Upon complexation of biotinylated TRAIL with RGDPEG-avidin, a bulky structure has been introduced onto the surface of the therapeutic protein. The assembled complexes, however, fully maintained their TRAIL activity indicating that steric hindrance did not interfere in the TRAIL-death receptor ligation. Figure 7 showed similar proapoptotic activities for either biotinylated TRAIL or the complexes formed in the presence of excess of RGDPEG-avidin. Remarkably, we observed an enhancement of caspase activity when the excess of RGDPEG-avidin was removed, leaving only surface-exposed RGDPEG-avidin at the endothelial cell layer. One of the explanations that may account for this is the assembly of multiple TRAIL molecules onto a single avidin, thereby affording a ‘mega’TRAIL complex with enhanced death-inducing activity, as was also observed for multimeric forms of Fas-ligand [29]. Such types of complexes would facilitate the close assembly of multiple death receptors, thereby potentiating the induction of death signaling. The presence of an excess of RGDPEG-avidin would avoid the formation of such complexes, as both components were added in a 100:1 molar ratio of avidin:TRAIL. Gently washing of the endothelial cells after RGDPEG-avidin incubation  removed the excess of unbound RGDPEG-avidin, leaving only cell-associated avidin after incubation at room termperature. These conditions therefore provided an excess of biotinylated TRAIL over avidin, now favoring the formation of multimeric TRAIL complexes. Alternatively, the exposure of TRAIL via RGD-PEG-avidin onto the surface of endothelial cells may explain its increased activity, as has been observed for chimeric TRAIL variants directed to tumor receptors [10].
HUVEC were not killed by TRAIL or its derivatives, as has been described by others [30]. On the other hand, although TRAIL does not induce apoptosis in endothelial cells, it activates other processes such as production of NO and reactive oxygen species which may alter the tumor responsiveness to TRAIL [31]. This can  not explain the results shown in Fig. 7, since endothelial cells were preincubated with RGDPEG-avidin only and preincubations were performed at room temperature rather than at 37°C. Such effects of RGD-equipped TRAIL may occur however in vivo. In addition, tumor derived growth factors might sensitize endothelial cells to TRAIL’s apoptosis inducing capacity, similar to the findings reported for TNF-sensitivity [32].
The complexation of biotinylated TRAIL with RGDPEG-avidin and its association with angiogenic endothelial cells may afford an increased accumulation into the tumor blood vessels, similar to the enhanced tumor accumulation of other RGD-modified proteins [18]. An unresolved issue is whether RGD-avidin/TRAIL complexes bound to the tumor vasculature can interact with tumor cells behind the endothelial lining in vivo. To interact with tumor cells and induce tumor apoptosis, TRAIL complexes bound to the tumor vasculature should dissociate and penetrate into the tumor tissue. Several considerations may be important in this respect, such as the expression of αvβ3 integrin on both luminal and abluminal side of the endothelium and the fact that many tumor cells express αvβ3 integrin [18]. Both of these will favor the interaction of RGD-avidin complexed TRAIL with tumor cells. Furthermore, RGD-avidin binding to endothelial cells is less tight than the typical antibody binding. This may favor redistribution of the complexes upon their initial binding to αvβ3 integrin. Homing to the tumor vasculature will provide elevated concentrations within the tumor microenvironment, which may subsequently provide elevated intratumoral concentrations. Together with the observed enhancement of TRAIL activity, this may favor a more efficient induction of apoptosis in tumor cells.
To summarize, we have prepared a novel type of RGD-targeted TRAIL with binding specificity for angiogenic endothelium and maintenance of strong cytotoxic activity. Whether in vivo application of this product  will lead to an enhanced tumor accumulation still needs to be investigated. We now succeeded in coupling the well known homing device RGD to the TRAIL molecule, without losing the apoptosis-inducing activity of TRAIL or the receptor-binding properties of RDG peptides. An enhanced activity could even be demonstrated in vitro. This may, in combination with an improved pharmacokinetic profile induced by the PEG and RGD moieties, lead to improved effects of TRAIL in vivo. Lastly, the followed synthetic approach offers opportunities for the complexation with alternative targeting ligands that bind directly to tumor cells, such as tumor-specific antibodies or peptide ligands.

apoptosis
therapeutic proteins
angiogenesis
drug targeting