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Apple Derived Cellulose Scaffolds for 3D Mammalian Cell Culture
Daniel J. Modulevsky,
Affiliations Centre for Interdisciplinary NanoPhysics, University of Ottawa, Ottawa, Ontario, Canada, Department of Biology, University of Ottawa, Ottawa, Ontario, Canada
Cory Lefebvre,
Affiliation Centre for Interdisciplinary NanoPhysics, University of Ottawa, Ottawa, Ontario, Canada
Kristina Haase,
Affiliations Centre for Interdisciplinary NanoPhysics, University of Ottawa, Ottawa, Ontario, Canada, Department of Physics, University of Ottawa, Ottawa, Ontario, Canada
Zeinab Al-Rekabi,
Andrew E. Pelling
* E-mail: a@pellinglab.net
Affiliations Centre for Interdisciplinary NanoPhysics, University of Ottawa, Ottawa, Ontario, Canada, Department of Biology, University of Ottawa, Ottawa, Ontario, Canada, Department of Physics, University of Ottawa, Ottawa, Ontario, Canada, Institute for Science, Society and Policy, University of Ottawa, Ottawa, Ontario, Canada
Daniel J. Modulevsky Cory Lefebvre ... Andrew E. Pelling
There are numerous approaches for producing natural and synthetic 3D scaffolds that support the proliferation of mammalian cells. 3D scaffolds better represent the natural cellular microenvironment and have many potential applications in vitro and in vivo. Here, we demonstrate that 3D cellulose scaffolds produced by decellularizing apple hypanthium tissue can be employed for in vitro 3D culture of NIH3T3 fibroblasts, mouse C2C12 muscle myoblasts and human HeLa epithelial cells. We show that these cells can adhere, invade and proliferate in the cellulose scaffolds. In addition, biochemical functionalization or chemical cross-linking can be employed to control the surface biochemistry and/or mechanical properties of the scaffold. The cells retain high viability even after 12 continuous weeks of culture and can achieve cell densities comparable with other natural and synthetic scaffold materials. Apple derived cellulose scaffolds are easily produced, inexpensive and originate from a renewable source. Taken together, these results demonstrate that naturally derived cellulose scaffolds offer a complementary approach to existing techniques for the in vitro culture of mammalian cells in a 3D environment.
Citation: Modulevsky DJ, Lefebvre C, Haase K, Al-Rekabi Z, Pelling AE (2014) Apple Derived Cellulose Scaffolds for 3D Mammalian Cell Culture. PLoS ONE 9(5): e97835. https://doi.org/10.1371/journal.pone.0097835
Editor: Irina Kerkis, Instituto Butantan, Brazil
Received: November 13, 2013; Accepted: April 24, 2014; Published: May 19, 2014
Copyright: © 2014 Modulevsky et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was made possible by support from the University of Ottawa and a Natural Science and Engineering Research Council (NSERC) Discovery Grant. C.L. was supported by an NSERC Undergraduate Student Research Award, Z.A.R. was supported by a graduate fellowship from the NSERC-CREATE Training Program in Quantitative. A.E.P. acknowledges generous support from the Canada Research Chairs Program and a Province of Ontario Early Researcher Award. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: Andrew Pelling is a PLOS ONE editorial board member and this does not alter the authors' adherence to all the PLOS ONE policies on sharing data and materials.
Development of novel biomaterials for the in vitro culture of cells in three-dimensional (3D) microenvironments has gained traction in recent years [1]–[6]. The motivation behind this development is to compensate for limitations of current two-dimensional (2D) cell culture practices. In particular, 2D plastic or glass substrates are ubiquitously employed to study many biological processes, despite the obvious structural and mechanical differences with the in vivo microenvironment. In vivo, cells are found in a complex extracellular matrix (ECM) whose biochemical and physical properties have a significant impact on numerous critical physiological and pathological processes [7]. Significant morphological and biological differences have already been observed between cells grown on 2D versus 3D microenvironments [8], [9]. It is has been routinely observed that primary cells isolated from tissues will become progressively flatter when cultured on conventional 2D surfaces [10], [11]. Conversely, cells cultured on 2D surfaces can regain their 3D morphologies when placed into a 3D culture scaffold [12]. 3D cell culture promises to more closely reflect the biochemical and physical properties of the cellular microenvironment found in tissues and organs [13], and so the development of novel biomaterials towards this effort is of considerable importance.
Both synthetic and naturally derived materials are currently employed in 3D culture methods, in order to create tuneable scaffolds engineered with specific biochemical and physical properties. Cellulose, the major component of plant cell walls, is an organic polysaccharide made of D-glucose subunits through β(1–4) bonds. Unlike the polysaccharides starch and glycogen, cellulose provides very little nutritional energy as the β(1–4) glycosidic bonds are difficult to digest and can only be broken down by cellulase [14]. As such, there has a great focus on using cellulose as a candidate biomaterial [12], [15]–[23]. Cellulose has previously been employed as a permeable dialysis membrane and as diffusion limiting membranes within biosensors[24]. As well, previous studies found that cellulose produced by bacteria could support the proliferation of mammalian cells [20], [25], [26]. Synthetically produced cellulose scaffolds have also been employed for 3D mammalian cell culture [2], [19], [21], [23]. Myocytes cultured on these synthetic cellulose scaffolds contained periodic myofibrils, a distinct cytoarchitectural element within mature cardiac myocytes [27]. As well, enhanced connectivity, in the form of increased gap junction density, and electrochemical connectivity, resulted from 3D culture, in comparison to cells grown on glass [27]. These examples suggest that cellulose may be a suitable material to support 3D cell growth. Moreover, cellulose is widely available as it is the most common organic polymer, accounting for 1.5×1012 tons of total annual biomass production [28].
Apple hypanthium tissue has an internal structure composed of cell walls that encompass pores and air pockets, facilitating the transport of nutrients and water throughout the fleshy tissue. These naturally developed characteristics, are important in any scaffold employed for 3D cell culture [29]. In order to act as a 3D scaffold, the apple hypanthium tissue must first be decellularized in order to remove existing nucleic acids, lipids, and proteins, producing a purified cellulose scaffold. Decellularization is now commonly employed on mammalian tissues to selectively remove cellular components while leaving behind an intact ECM [30]–[33] Typically, mammalian tissues are bathed in solutions, detergents and/or proteases, in order to produce a decellularized matrix that retains the shape of the original tissue[34]–[43]. Decellularized tissue can then be repopulated with new cells, in order to produce new functional organs. Hearts, kidneys have been decellularized and reseeded with various cells [33], [41]–[43]. As well, functional bladders and lungs have be produced and transplanted into animals using this technique [44], [45]. Importantly, decellularized tissue also maintains a well conserved native ECM architecture and cell-ECM binding domains [41].
In this study, we hypothesized that decellularized apple hypanthium tissue might provide an easily produced scaffold for 3D cell culture. The major aim of this study was to demonstrate that mammalian cells would successfully proliferate within a 3D cellulose scaffold in vitro. Through modification of an existing decellularization protocol, we generated apple-derived cellulose scaffolds for cell culture. We examined how three mammalian cell types (mouse NIH3T3 fibroblasts, mouse C2C12 myoblasts and human HeLa epithelial cells) proliferated within these scaffolds, for up to twelve weeks. Phase contrast microscopy, laser scanning confocal microscopy and scanning electron microscopy were used to characterize the structure of the scaffolds, cell growth, cell morphology and the influence of the scaffolds on the actin cytoskeleton. We also modified the surface biochemistry and mechanical properties of the cellulose scaffolds by collagen functionalization, or chemical cross-linking with glutaraldehyde. Atomic force microscopy was employed to quantify the effect of these modifications on the mechanical properties of the scaffolds. We demonstrate that the 3 mammalian cell lines used in this study were able to proliferate and remain viable in the 3D cellulose scaffold in vitro, achieving cell densities similar to other synthetic and natural biomaterials. Given the natural porosity and ease of production of cellulose scaffolds, as well as the ability to modify their mechanical properties, we demonstrate that cellulose scaffolds are a potentially useful biomaterial that can be successfully employed for in vitro 3D cell culture.
Apple tissue preparation, decellularization and storage
McIntosh Red apples (Canada Fancy) were stored at 4°C in the dark for a maximum of two weeks. In order to prepare apple sections, the fruit was first chilled in a −20°C freezer for 5 minutes prior to being cut with a mandolin slicer to a uniform thickness of 1.20±0.14 mm, measured with a vernier caliper (Fig. 1 A-B). Only the outer (hypanthium) tissue of the apple was used. Slices containing visible ovary-core tissue were not used. The slices were then cut into 2.0×0.5 cm segments parallel to the direction of the apple pedicel (Fig. 1 C). Apple tissue was then decellularized by using a well-established protocol [41] for removing cellular material and DNA from tissue samples while leaving behind an intact and three-dimensional scaffold. Individual apple tissue samples were placed in sterilized 2.5 mL microcentrifuge tubes and 2 mL of 0.5% sodium dodecyl sulphate (SDS) (Sigma-Aldrich) solution was added to each tube. Samples were shaken for 12 hours at 160 RPM at room temperature (Fig. 1 D). The resultant cellulose scaffolds were then transferred into new sterile microcentrifuge tubes, washed and incubated for 6 hours in PBS (Sigma-Aldrich) with 1% streptomycin/penicillin (HyClone) and 1% amphotericin B (Wisent). At this point, the samples were immediately used or stored in PBS at 4°C for no more than 2 week.
Figure 1. A cartoon schematic representing the apple tissue decellularization and mammalian cell seeding protocol used in this study.
A) McIntosh Red apples were exposed to −20°C temperatures for a max duration of 5 minutes, to increase the firmness of the outer apple hypanthium tissue. B) Uniform 1.2±0.1 mm thick slices of the apples were obtained using a mandolin slicer. Slices containing any of the ovary core of the apple were removed. C) The apple slices were cut into uniform 2.0 by 0.5 cm segments that were placed in individual microcentrifuge tubes. D) A 0.5% SDS solution was added to the microcentrifuge tubes and placed on a shaker for 12 hours at room temperature. The scaffolds were then rinsed repeatedly with PBS and allowed to incubate in a PBS solution with 1% streptomycin/penicillin and 1% amphotericin B for 6 hours at room temperature. E) The scaffolds were then coated with Type 1 collagen, chemically cross linked with glutaraldehyde or incubated in PBS. F) All the samples were then incubated in mammalian cell culture medium (DMEM) for 12 hours in a standard tissue culture incubator maintained at 37°C and 5% CO2. G) The scaffolds were placed in PDMS coated 24 well plates and a 40 µL cell suspension was placed on each. After 6 hours in the incubator the wells were filled with DMEM and cells cultured for up to 12 weeks.
Post-decellularization treatments
Here, we examined cell proliferation and invasion into native, collagen functionalized, or chemically cross-linked cellulose scaffolds. In order to functionalize scaffolds with collagen, samples were incubated for 6 hours in a solution of 10% acetic acid and 1 µg/mL rat tail collagen type I (Invitrogen), followed by washing in PBS before use. To chemically cross-link the scaffolds, the samples were incubated in a 1% EM-grade glutaraldehyde solution (Sigma-Aldrich) for 6 hours. The scaffolds were then rinsed in PBS and incubated in a solution of 1% sodium borohydride (Sigma-Aldrich) overnight in order to reduce any unreacted glutaraldehyde (Fig. 1E). Prior to seeding cells into the scaffolds, all samples (native, collagen coated, or cross-linked) were incubated in mammalian cell culture medium (described below) for 12 hours in a standard tissue culture incubator maintained at 37°C with 5% CO2 (Fig. 1F).
C2C12 mouse myoblasts, NIH3T3 mouse fibroblasts and HeLa human epithelial cell lines were used in this study (all obtained from the American Type Culture Collection (ATCC)). Cells were cultured in standard cell culture media (high glucose DMEM (HyClone), supplemented with 10% fetal bovine serum (HyClone), 1% penicillin/streptomycin (HyClone) and 1% amphotericin B (Wisent) at 37°C and 5% CO2 in T75 flasks (Thermo Scientific). Culture media was exchanged every second day and the cells were passaged at 80% confluence.
In vitro cell culture in cellulose scaffolds
The scaffold seeding procedure took place in 24-well tissue culture plates. Each well was individually coated with polydimethylisiloxane (PDMS) to create a hydrophobic surface in order to prevent the adhesion of cells. A 1:10 solution of curing agent: elastomer (Sylgard 184, Ellsworth Adhesives) was poured into each well. The PDMS was cured for 2 hours at 80°C, and was allowed to cool to room temperature, then rinsed with PBS. Scaffolds were cut into 0.5×0.5 cm pieces and placed within each well. A 40 µL droplet containing 6×106 cells was carefully formed on top of each scaffold. The samples were placed in the incubator for 6 hours to allow the cells to adhere to the scaffolds. Subsequently, 2 mL of DMEM was added to each well and the samples were incubated for 48 hours. At this point, samples containing mammalian cells were then carefully transferred into new 24-well PDMS-coated tissue culture plates. For continued cell proliferation, the culture media was exchanged every day and scaffolds were moved into new 24-well plates every 2 weeks.
Immunofluorescence staining
The actin cytoskeleton and nucleus of mammalian cells, cultured on glass or within the scaffolds, were stained according to previous protocols [46], [47]. Briefly, samples were fixed with 3.5% paraformaldehyde and permeabilized with Triton X-100 at 37°C. Actin was stained with phalloidin conjugated to Alexa Fluor 488 (Invitrogen) and nuclei were stained by labelling the DNA with DAPI (Invitrogen). Samples were then mounted in Vectashield (Vector Labs).
In order to simultaneously stain the cellulose scaffold and mammalian cells, we first fixed the samples as described above, and then washed them with PBS 3 times. To label the apple cell walls, we used an established protocol described previously by Trueunit et al. (2008) [48]. The samples were rinsed with water and incubated in 1% periodic acid (Sigma-Aldrich) at room temperature for 40 minutes. The tissue was rinsed again with water and incubated in Schiff reagent (100 mM sodium metabisulphite and 0.15 N HCl) with 100 mg/mL propidium iodide (Invitrogen) for 2 hours. The samples were then washed with PBS. To visualize the mammalian cells within the apple tissue, the samples were incubated with a solution of 5 µg/mL wheat germ agglutinin (WGA) 488 (Invitrogen) and 1 µg/mL Hoechst 33342 (Invitrogen) in HBSS (20 mM HEPES at pH 7.4; 120 mM NaCl; 5.3 mM KCl; 0.8 mM MgSO4; 1.8 mM CaCl2; and 11.1 mM dextrose). WGA and Hoechst 33342 are live cell dyes that label the mammalian cell membrane and nucleus, respectively. The samples were then transferred onto microscope slides and mounted in a chloral hydrate solution (4 g chloral hydrate, 1 mL glycerol, and 2 mL water). Slides were kept overnight at room temperature in a closed environment to prevent dehydration. The samples were then placed in PBS until ready for imaging.
We also labelled samples to test for long-term mammalian cell viability. In these cases, cells were maintained in culture for 12 weeks and then stained with a solution of 1 µg/mL Hoechst 33342, which stains the nuclei of all cells, and 1µg/mL Propidium iodide (PI), which is cell membrane impermeable and will only stain the nucleic acids of apoptotic or necrotic cells. Samples were then fixed with 3.5% paraformaldehyde as above and then submerged in PBS until ready for confocal imaging. In order to quantify the number of viable cells we prepared and stained n = 3 samples. Individual Hoechst-positive and PI-positive cells were automatically counted using the particle analyzer function on ImageJ.
Optical Microscopy
Confocal imaging was performed on an A1R high speed laser scanning confocal system on a TiE inverted optical microscope platform (Nikon, Canada) with appropriate laser lines and filter sets. Transmitted light images were acquired on an inverted TiE microscope (Nikon, Canada) with phase contrast optics. Images were analyzed using ImageJ open access software (http://rsbweb.nih.gov/ij/). Brightness and contrast adjustments were the only manipulations performed to images.
Scaffolds containing mammalian cells were first fixed with 3.5% paraformaldehyde as presented above, and then gently washed repeatedly with PBS. The samples were then dehydrated through successive gradients of ethanol (50%, 70%, 95% and 100%) and dried within a lyophilizer. Samples were then gold-coated at a current of 15 mA for 3 minutes with a Hitachi E-1010 ion sputter device. SEM imaging was conducted at voltages ranging from 2.00–10.0 kV on a JEOL JSM-7500F FESEM.
Atomic force microscopy (AFM)
AFM was employed to measure the mechanical properties of the native, collagen functionalized, or chemically cross-linked (n = 3 in each case) cellulose scaffolds. In all cases, tip-less PNP-TR-TL (Nano world) AFM cantilevers were modified with a 10 µm polystyrene bead (Fluka) using an optical adhesive (Norland), cured in a UV cross-linker (Spectroline Select Series). Cantilevers possessed an average spring constant of 37±5 pN/nm as determined using the thermal fluctuation method [49], [50]. Local mechanical properties were measured by 5–15 force-indentation curves collected at 10–15 randomly chosen locations at a rate of 1 Hz. A total of n = 200 measurements were acquired for each sample. PUNIAS software was used to fit the first 200 nm of indentation to the Hertz model for a spherical indenter, using a Poisson ratio of 0.5 [46], [50], [51].
All presented values are the average ± standard deviation. Where applicable, we assessed statistical significance by performing a two-sample student's t-test (α<0.05).
Preparation of cellulose scaffolds
As described in the Materials and Methods section, apple hypanthium tissue was cut to uniform size and decellularized following established protocols (Fig. 1) [41]. Hypanthium tissue was employed as it is rich in cellulose and contains very few cells [52], [53] Decellularization protocols were employed to ensure the complete removal of any remaining plant cells, nucleic acids and biomacromolecules. After processing the samples, a highly porous structure is observed with phase contrast microscopy (Fig. 2A). The apple tissue has evolved as a very porous structure, with cell wall cavities observed throughout the sample, allowing for facilitated nutrient transfer throughout. Cellulose scaffolds were then fixed and dehydrated for SEM imaging. Samples were cut horizontally down the mid-section revealing the interior surface. A highly porous and relatively robust scaffold is clearly observed (Fig. 2B). In all cases, the cellulose scaffold was the only apparent feature observed in all images, as no other identifiable structures were witnessed (i.e. cellular organelles or otherwise).
Figure 2. Decellularized cellulose scaffolds.
A) Phase contrast image of cellulose cell wall structure in a decellularized apple tissue sample. The dark lines correspond to distinct cellulose structures which form a three dimensional matrix. B) SEM image of a similar cellulose scaffold revealing its three dimensional nature and large cavities. Scale bar = 200 µm.
Mechanical properties of native and modified cellulose scaffolds
We employed two post-decellularization functionalization protocols in order to examine the ease of modification of the mechanical properties of these cellulose scaffolds. The two modifications included functionalization of the scaffold with type I collagen, or chemically cross-linking of the scaffold with glutaraldehyde. These modifications allowed us to control the biochemical environment of the scaffolds, and alter their mechanical properties. AFM was used to quantify the local elasticity of the scaffolds in response to each treatment. We measured the elasticity of four specific samples: untreated (native), decellularized (SDS), decellularized and collagen functionalized (SDS+Coll), and finally decellularized and glutaraldehyde cross-linked (SDS+GA) tissue. Native tissue, SDS, SDS+Coll and SDS+GA scaffolds possessed an elasticity of 0.9±0.1 kPa, 1.1±0.1 kPa, 2.2±0.2 kPa and 4.1±0.3 kPa, respectively (Fig. 3A). The native and SDS scaffolds did not display any significant difference in mechanical properties (p>0.05). Both the SDS+Coll and SDS+GA scaffolds displayed a significant increase in elasticity compared to the native and decellularized scaffolds (p<0.001). These results demonstrate that the local elasticity of the scaffolds can clearly be controlled to fall within a range which mimics some mammalian tissues [54], [55] Finally, C2C12 cells were seeded into different scaffold preparations (SDS, SDS+Coll and SDS+GA, n = 3 in each case) and cultured for 2 weeks (Fig 3B-D). Phase contrast microscopy revealed the presence of mammalian cells in each of the scaffolds compared to the cellulose scaffold presented in Fig. 2A. The images shown are representative of the n = 3 scaffolds prepared in each case.
Figure 3. The mechanical properties of functionalized cellulose 3D scaffolds and C2C12 myoblast cultured within the 3D cellulose scaffolds.
A) The local mechanical elasticity of native tissue, decellularized (SDS), collagen functionalized (SDS+Coll) and glutaraldehyde (SDS+GA) cross-linked cellulose scaffolds. The native tissue and unmodified scaffolds do not display any significant difference in mechanical properties. Both the collagen functionalized and chemically cross-linked scaffolds displayed a significant increase in elasticity compared to the DMEM scaffolds (*** = p<0.001). The (B) decellularized (SDS), (C) Collagen functionalized (SDS+Coll) and (D) glutaraldheyde cross-linked (SDS+GA) scaffolds all support the growth of C2C12 cells. Phase contrast images of C2C12 cells after two weeks of growth reveal the presence of cell colonies. Scale bar = 200 µm.
Mammalian cell culture in native, collagen functionalized, and chemically cross-linked cellulose scaffolds
To test the ability of the cellulose scaffolds for supporting 3D mammalian cell culture, we examined the proliferation of mouse C2C12 myoblasts, mouse NIH3T3 fibroblasts and human HeLa epithelial cells within the native, collagen functionalized, or chemically cross-linked cellulose scaffolds (n = 3 for each case). In all cases, proliferation of cells was similar. After 4 weeks in culture, immunofluorescent images were generated for every sample, to selectively visualize the mammalian cells within the apple cell wall. A distinct cell wall structure was observed for all conditions, seen as a uniform thin red structure forming individual cavities of consistent size (see Fig. 4). Interestingly, similar observations were made of the cellulose structure in the SEM images (Fig. 2b). The cell membranes (green) and nucleus (blue) of C2C12 myoblasts (Fig. 4A), mouse NIH3T3 fibroblasts (Fig. 4B) and human HeLa epithelial cells (Fig. 4C) clearly proliferate very well in unmodified (Fig. 4A-C), as well as modified scaffolds (data not shown). Cells were observed to be growing on the surface of the scaffolds.
Figure 4. Fixed and stained NIH3T3, C2C12 and HeLa cells cultured on 3D cellulose scaffolds.
Specific fluorescent staining of (A) NIH3T3, (B) C2C12 and (C) HeLa mammalian cells within the cellulose scaffolds and subsequent laser scanning confocal microscopy reveals the cellulose structure (red), mammalian cell membranes (green) and nuclei (blue). Cells were cultured in these scaffolds for four weeks prior to staining and imaging. Confocal volumes were acquired and projected in the XY and ZY plane. The ZY orthogonal views demonstrate the depth of cell proliferation within the cellulose scaffold. The top and bottom surfaces of the scaffold are indicated. Scale bars: XY = 300 µm, ZY = 100 µm. D) SEM image of a cellulose scaffold cross section after being seeded with C2C12 cells that were allowed to proliferate for four weeks. The cells were digitally colourized in order to increase contrast between the cells and cellulose structure (Scale bar: 50 µm)
It was also observed that the majority of cells are in closely interacting aggregates within the individual cell wall cavities. To further examine the proliferation within the cellulose structure, orthogonal views of volume-images were generated. The green (membrane) and blue (nucleus) signals can be seen within the interior of the cellulose structure, thus demonstrating that the cells proliferate deep within the scaffolds. Fluorescent signals can be seen up to depths of approximately 120 µm from the surface of the scaffolds. It is important to note that the loss of fluorescence signal coincides with the limit of imaging depth of the confocal microscope. Hence, the actual degree of cell invasion into the scaffold cannot be determined with confocal microscopy alone. To further investigate the degree of cellular invasion into the cellulose scaffolds we fixed and dehydrated (n = 3) native scaffolds for SEM imaging. The samples were prepared in a way that revealed the interior surface of the cellulose scaffolds. SEM images also reveal that cells can migrate within the scaffolds. In Fig 4D, we present images of C2C12 cells that have migrated towards the interior of the sample. Cells are clearly visible within the scaffold (Fig. 4D) in comparison to SEM of an un-seeded scaffold (Fig. 2B). Cells in the scaffold can be seen attached to the cellulose structure, with their morphologies varying between round and spread, consistent with other mammalian cells grown in other natural and synthetic 3D scaffolds [56]–[58].
Finally, the morphology of the actin cytoskeleton was also examined for all cell types in unmodified cellulose scaffolds (n = 3 for each cell type). In this case, cells were only cultured for 1 week, as the actin cytoskeleton was more easily resolved in lower density samples. All mammalian cell types within the scaffolds displayed clear actin stress fibers, consistent with strongly attached cells, as those demonstrated on solid 2D substrates [46], [47], [59] (Fig. 5). These results demonstrate that the mammalian cells were well-adhered to the cellulose scaffold, rather than simply being physically confined within the scaffold.
Figure 5. Fixed and stained images of cells actin cytoskeleton cultured within the 3D cellulose scaffold.
A) NIH3T3, B) C2C12 and C) HeLa cells were cultured onto the cellulose scaffolds for 2 weeks prior to stained for actin (green) and cell nuclei (blue). (. NIH3T3 and C2C12 cells display characteristic actin stress fibres found in cultured cells. HeLa cells also display characteristic actin structures including fewer prominent stress fibres and a large amount of cortical actin localization. Scale bar = 25 µm and applies to all.
Proliferation and Viability
In order to quantitatively assess long-term cell proliferation and viability, confocal images were acquired throughout the cellulose scaffold after 1, 8 and 12 weeks of continuous cell culture. To assess proliferation, confocal volumes of cell nuclei were acquired at random locations on n = 3 scaffolds seeded with NIH3T3, C2C12 or HeLa cells. For each cell type, n = 3 randomly chosen 1.6×106 µm2 imaging regions were chosen, and confocal imaging was performed. The imaging depth required to capture all cells in the sample area was observed to change due to the fact that cells were invading deeper into the scaffold over time. Using this data, cells were counted and shown to increase in number by several-fold over a 12-week period (Fig. 6A). HeLa and C2C12 cells were observed to increase in number at a similar rate, which was higher than NIH3T3 cells. Regardless, all cell types exhibited a three- to four-fold increase in number over a 12-week period of continuous culture.
Figure 6. Cell proliferation and viability over time.
A) NIH3T3, C2C12 and HeLa cells were cultured individually in cellulose n = 3 scaffolds for 1, 8 and 2 weeks and then imaged with confocal microscopy after being stained with Hoechst 33342. Cells were counted at each time point and an increase in cell population is clearly observed. B) After 12 weeks of culture, C2C12 cells were fixed and stained with Hoechst 33342 (blue: viable cells) and Propidium iodide (PI) (red: apoptotic/necrotic cells). Confocal volumes were acquired and projected in the XY and ZY plane and reveal that cells have proliferated throughout the structure during the 12-week culture. The cells that are apoptotic/necrotic are found in deeper regions of the scaffold. The top and bottom surfaces of the scaffold are indicated. The number of live (Hoechst(+)) and dead (Hoechst/PI(+)) cells were counted and it was found that ∼98% of the cells within the scaffold are viable. Data is shown for C2C12 cells, but is similar for NIH3T3 and HeLa cells (data not shown). Scale bar: B = 200 µm for XY and 100 µm for ZY.
We also examined cell viability within these cellulose scaffolds after 12-weeks of continuous culture. Scaffolds (n = 3) were incubated with Propidium iodide and Hoechst 33342 to specifically label apoptotic/necrotic or living cells, respectively (Fig. 6B). In all cases, Hoechst labelled all cell nuclei (live and dead) throughout the sample. Conversely, Propidium iodide specifically labelled a minority of dead cells, which were mainly located towards the interior portion of the scaffold. Cell viability was quantified by counting the Hoescht-positive (live) and PI/Hoescht-positive (dead) nuclei of cells. The majority of cells (98±1% of C2C12 cells) cultured on the apple scaffold were Hoechst-positive after 12 weeks in culture (Figure 6C), with identical results in the other cell types (data not shown). These results demonstrated a high degree of viability for long-term cultures on the surfaces of these cellulose scaffolds. Orthogonal projections demonstrated that a small population of apoptotic/necrotic cells were located approximately 120 µm towards the interior of the cellulose scaffold (see Fig. 6B). Even in regions of increased cell density there was sufficient porosity within the cellulose scaffold to allow for media/nutrient transfer to the interior of the scaffolds.
Cells in a 3D environment, whether an artificial extracellular matrix or living tissues, often display numerous morphological and biochemical differences compared to cells cultured on a 2D surface [8]. These differences are under intense scrutiny, as many studies have demonstrated the importance of spatial cues within the ECM. For example, geometry and special cues have been shown to affect cell morphology, mRNA signalling, and differentiation [5], [9], [60]–[62]. Therefore, the use of 3D scaffolds for in vitro cell culture is beneficial towards our fundamental understanding of cell biology, and has consequently has many potential applications. The development and use of 3D biocompatible scaffolds does however present several challenges. Biomaterials must be biocompatible, biodegradable, allow for surface modifications, and be cost effective. Two methods of production have emerged. The first of these methods, is the use of artificial scaffolds synthesized from (bio)polymers. Artificial scaffolds have the advantage of allowing for exceptional control over the material, allowing for tuning of biochemical and structural properties of the scaffold [63]–[67]. On the other hand, decellularization has been employed to produce natural 3D scaffolds from existing tissue [13], [33], [44], [65]–[68]. Decellularization employs various reagents to lyse and remove cells from the ECM of a given tissue sample [32], [34]. Though this approach lacks fine control over the physical and biochemical properties of the scaffold, it results in an easily obtained, naturally derived, scaffold that has been employed in the creation of functional organs [33], [41]–[44].
Herein, decellularized apple hypanthium tissue is shown to provide several characteristics that are desirable for producing 3D cellular scaffolds. Apple hypanthium is primarily composed of plant cell walls which form a porous network of cavities in which the plant cells reside [69]. This porous environment allows for facilitated transfer of nutrients throughout the tissue. In addition, this environment is also ideal for decellularization as it allows for rapid exchange of detergents, buffers and media without the use of perfusion systems. We were able to rapidly produce decellularized cellulose scaffolds by simply using a detergent solution. Importantly, mouse fibroblast (NIH3T3), mouse muscle myoblast (C2C12) and human epithelial (HeLa) cells were all able to completely invade, proliferate and remain viable after a long term period of culture (12 weeks) within the cellulose scaffolds.
The mechanical and biochemical properties of the ECM are known to play a very important role in governing cellular physiology [5], [60]–[62]. The apple hypanthium-derived scaffolds, used in this study, are composed of cellulose that is assembled into microfibrils which are then cross-linked by hemicellulose [70], and resemble the porous nature of the ECM. Here, using AFM, we demonstrated that the local Young's modulus of the scaffold could be altered through chemical cross-linking or biochemical functionalization. We found that the native and SDS samples displayed no statistical difference in local Young's modulus (p<0.05), demonstrating that SDS treatment did not alter the mechanics of the cellulose structure [31]. Furthermore, it was also observed that scaffolds functionalized with collagen or cross-linked with glutaraldehyde displayed a significant two to four-fold increase in Young's modulus (p<0.001) compared to the native tissue or unfunctionalized decellularized scaffolds. Our results are consistent with previous studies, which demonstrate that collagen functionalization or glutaraldehyde cross-linking tends to increase the Young's modulus of biological materials [71]–[73]. The ability to tune the cellulose scaffold rigidity is critical as it has been long established that cells preferred matrix with specific rigidities to differentiate and acquire their specific cellular properties [32], [34], [60], [69], [70]. Hence, we showed that it is possible to control and modulate the mechanical rigidity of the cellulose scaffold.
Previous studies employing mechanical compression testing of bulk apple samples have reported significantly higher Young's modulus measurements, in the MPa range [74], [75], compared to the ones obtained in our study. Their technical approaches measure an elasticity that reflects the forces required to deform all of the cells and cellulose structures within a large region of apple tissue. In contrast, we employed an AFM with a modified cantilever probe using a 10 µm colloid polystyrene bead in order to probe local elasticities. Young's modulus measurements obtained with this approach are reflective of the local deformation of a small microscale region of the cellulose scaffold. Due to the local nature of AFM measurements, the measured Young's modulus more accurately reflects the mechanical properties experienced by the cells. Interestingly, in a recent study, the local Young's modulus of tomato plant tissue was determined to be in the same order of magnitude of our measurements, ∼7–29 kPa [76]. These results were also obtained using AFM and a modified cantilever with an attached 10 µm colloid. Importantly however, under all conditions and for differing Young's moduli, cells were observed to rapidly grow and proliferate within the cellulose scaffolds.
Employing a staining protocol established by Truernit et al. [48], we simultaneously visualized the cellulose structure along with the embedded cells. Confocal microscopy images revealed that all three cell types used in this study were found on the surfaces and the interior cavities within the cellulose scaffold. Cells were reliably imaged to a depth of approximately 120 µm below the upper surface of the scaffold. In all three cell types we also observed the presence of distinct F-actin stress fibres, which are a morphological characteristic of substrate-adhered cells [47], [77], [78]. Both fibroblasts and myoblasts cultured in our scaffolds possessed pronounced actin stress fibres, similar to those found in the same cells cultured on 2D substrates [46], [47]. HeLa cells were observed to express cortical actin and actin stress fibres. In this case, stress fibres were less prominent than those observed within the fibroblasts and myoblasts, however the observed morphology of the actin cytoskeleton is very consistent with HeLa cells cultured on 2D substrates [78]. The results clearly demonstrate that all cell types studied here were able to proliferate, displaying strong attachment to the surfaces of the cellulose scaffolds.
A constraint of confocal microscopy is the limited depth at which a sample can be imaged. In order to determine whether the mammalian cells migrated towards the interior of the scaffold, the sample was fixed, frozen, and then fractured perpendicular to the top surface. This process allowed us to image the interior of the cellulose scaffold using SEM. In this case, C2C12 cells were cultured on the scaffolds and subsequently imaged. Cells were found deep within the interior of the scaffold and to possess morphologies that were both rounded, as well as flat and spread-out along the cellulose surfaces, analogous to that of myoblasts cells in other 3D environments [79], [80]. Interestingly, the cellulose scaffold appears in a collapsed/compressed state in the presence of mammalian cells. When mammalian cells are absent from the scaffold, such morphologies are not observed, confirming that the preparation for SEM imaging is not causing this effect. At present, the origin of this change in morphology is not well understood, however we hypothesize it may be due to the presence of cellular traction forces, adsorption of serum proteins on the scaffold, or cellular remodelling/degradation of the scaffold.
To quantify proliferation, we counted the number of cells present in the scaffolds after 1,8 and 12 weeks of continuous in vitro culture. In all cases, cells were observed to proliferate and increase in number by three to four-fold. However, it was observed that C2C12 and HeLa cells had nearly twice the number of NIH3T3 cells. This is an intriguing result as doubling rate of NIH3T3 cultured on glass (18 hours) is slightly lower than that of HeLa cells (24 hours) [81], [82]. Viability assays also reveal that a high percentage of cells remained viable even after 12 weeks of continuous in vitro culture. The majority of apoptotic cells were present towards the interior of the cellulose scaffold. This is likely caused by a lack of media turnover, leading to an insufficient supply of nutrients and oxygen, a common phenomenon in 3D culture models [83]. 3D spheroid cell cultures that often contain a necrotic core of cells due to an insufficient supply of nutrients and oxygen [83].
Although the results show that cells proliferate and remain viable in the scaffolds, their porosity can have an important influence on the observed results. The high porosity of the scaffolds offers both advantages and disadvantages. The high porosity of the scaffold allows cells to pass through the material without a high degree of confinement. This leads to a low seeding efficiency that can be problematic in some applications (the culture of rare cells, etc.). For example, in this study each scaffold (∼1 mm3 total volume) was initially seeded at a density of ∼6×106 cells/mm3. However, cell density was lower after 1-week of culture (∼2×106 cells/mm3) due to the loss of cells during media exchange. Therefore, a large amount of time (12-weeks) is required for the cells to slowly proliferate and invade the entire scaffold. This is not surprising given the issues highlighted above, and the fact that these mammalian cells are proliferating in a scaffold that is composed largely of cellulose as opposed to ECM proteins. Although there is a slow proliferation, the cell density does approach the levels reported in other biomaterials [41], [43], [84]. The efficiency of the initial seeding might be improved through further biochemical modification or chemical cross-linking of the scaffolds to increase the binding potential. Future work will investigate scaffold modification to control porosity and improve seeding efficiency. Alternatively, naturally occurring cellulose with lower porosity might be found in other apple species or plant types. Regardless, future studies will examine the use of higher density cellulose scaffolds to increase seeding efficiency, which will lower the time required to fully populate the scaffolds.
In addition, the high porosity of the scaffolds likely leads to an overestimation of cell viability. Dead cells can be easily washed away during media exchange, lowering the observed number of dead cells in the scaffold. Furthermore, the limited depth associated with confocal microscopy will also influence the ability to assay the physiological state of the cells deep within the scaffold. Other viability assays are also impeded by the nature of the cellulose scaffold. In particular, Trypan blue assay is problematic as the dye interacts strongly with the scaffold, which significantly impedes cell counting and quantification.
Although these limitations exist, mammalian cells are clearly able to proliferate, invade and possess an intact actin cytoskeleton within the scaffolds. In all three cases, cells were observed to interact closely with each other and the scaffold in large, long-range networks. Furthermore, theses cellulose scaffolds represent a very low-cost approach to studying the proliferation and invasion of cells in 3D. These cellulose biomaterials are highly complementary with the myriad of possibilities for the study of 3D cell biology in vitro. Cellulose scaffolds are porous enough to allow for efficient nutrient/gas exchange, biocompatible, easily functionalized and their mechanical properties can be controlled.
The results presented in this study naturally give rise to questions about the in vivo applicability of cellulose scaffolds. At this point, it is still too early to speculate, as there are many unknowns about the biocompatibility of these scaffolds in vivo. The immunological response and the long-term stability of implanted cellulose-based biomaterials are still being studied [22], [85]. We have not examined these issues in this study, as our objective was only to demonstrate the suitability of apple-derived cellulose scaffolds in supporting the in vitro culture of mammalian cells. Numerous approaches for producing 3D matrices that support the culture of mammalian cells are available [86], however, many of these products are proprietary, expensive, or require chemical synthesis. Plant derived cellulose scaffolds offer an alternative approach for 3D culture, offering the advantage of ease of production and modification, reduced cost, the ability to fabricate the cellulose into shapes specific to the user. Taken together, our results demonstrate that natural apple-derived cellulose scaffolds can be produced by employing common decellularization approaches and will support 3D cell culture. The evolved porosity of the apple hypanthium tissue facilitates the initial decellularization and provides critical media transfer allowing for long-term cell viability. Three mammalian cell types were observed to proliferate, migrate, and remain viable in the scaffolds for up to 12 weeks (maximum length of study). In addition, biochemical functionalization or chemical cross-linking can also be employed to control the surface biochemistry and/or mechanical properties of the scaffold. The advantages of these cellulose scaffolds make them one of several potential biomaterial candidates for in vitro 3D cell culture that are currently available.
The authors wish to thank Dr. Yun Liu for assistance with SEM imaging, Safi Sayeed for generously donating samples for this study and Tristan Matheson for assistance with figure preparation.
Conceived and designed the experiments: DM AEP. Performed the experiments: DM CL ZAR. Analyzed the data: DM AEP KH. Contributed reagents/materials/analysis tools: DM CL ZAR KH. Wrote the paper: DM CL ZAR KH AEP.
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Cellular structures and organelles
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US5845640A - Chemiluminescent sources for photodynamic therapy and photomedicine - Google Patents
Chemiluminescent sources for photodynamic therapy and photomedicine Download PDF
fluorescer
chemiluminescent
Nabil M. Lawandy
Spectra Science Corp
1997-09-15 Application filed by Spectra Science Corp filed Critical Spectra Science Corp
1997-09-15 Assigned to SPECTRA SCIENCE CORPORATION reassignment SPECTRA SCIENCE CORPORATION ASSIGNMENT OF ASSIGNORS INTEREST (SEE DOCUMENT FOR DETAILS). Assignors: LAWANDY, NABIL M.
A61N5/00—Radiation therapy
A61N5/06—Radiation therapy using light
A61N5/0601—Apparatus for use inside the body
A61K41/00—Medicinal preparations obtained by treating materials with wave energy or particle radiation; Therapies using these preparations
A61K41/0057—Photodynamic therapy with a photosensitizer, i.e. agent able to produce reactive oxygen species upon exposure to light or radiation, e.g. UV or visible light; photocleavage of nucleic acids with an agent
A61N5/0613—Apparatus adapted for a specific treatment
A61N5/062—Photodynamic therapy, i.e. excitation of an agent
F21K—NON-ELECTRIC LIGHT SOURCES USING LUMINESCENCE; LIGHT SOURCES USING ELECTROCHEMILUMINESCENCE; LIGHT SOURCES USING CHARGES OF COMBUSTIBLE MATERIAL; LIGHT SOURCES USING SEMICONDUCTOR DEVICES AS LIGHT-GENERATING ELEMENTS; LIGHT SOURCES NOT OTHERWISE PROVIDED FOR
F21K2/00—Non-electric light sources using luminescence; Light sources using electrochemiluminescence
F21K2/06—Non-electric light sources using luminescence; Light sources using electrochemiluminescence using chemiluminescence
A61N2005/0635—Radiation therapy using light characterised by the body area to be irradiated
A61N2005/0643—Applicators, probes irradiating specific body areas in close proximity
A61N2005/0645—Applicators worn by the patient
An optical source (10) for exciting a photosensitive drug that responds to light within a predetermined band of wavelengths. The optical source uses a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths. The chemiluminescent source can be flowed during use. In one embodiment the chemiluminescent source is stored in a heated reservoir, and is pumped from the reservoir through a conduit, such as a catheter. A patch can be used for spreading the chemiluminescent source over a wider area. An active feedback system can also be used for controlling the output light. In preferred embodiments a plurality of reactants are mixed for forming a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths. The mixing of the reactants causes an excitation of the at least one selected fluorescer by energy transfer from an energetic immediate in a chemiluminescent reaction to provide a fluorescent emission in the predetermined band of wavelengths.
This application claims the benefit of U.S. provisional application(s) No(s). 60/036,246, Jan. 24, 1997.
This invention relates generally to optically-based therapeutic procedures.
Photodynamic Therapy (PDT) uses specifically designed drugs such as FOSCAN (Scotia Pharmaceuticals), a photodynamic therapy related drug, ALA (DUSA) and Photofrin (QLT Phototherapeutics) to destroy rapidly dividing cells. These drugs are selectively retained or generated at rapidly dividing cells and are subsequently excited by light to produce the desired effects. The primary mode of activity usually involves energy transfer from these photoexcited drugs to O2 to produce superoxides or O2 in the singlet state. To date this excitation has been provided by lasers, lamps, and new materials such as LASERPRINT (a new material which provides laser action in amplifying scattering media). Some of these sources are expensive and require complicated delivery systems.
OBJECTS OF THE INVENTION
It is an object of this invention to provide an improved method and apparatus to excite an optically-responsive or photosensitive therapeutic drug.
It is a further object of this invention to provide an improved method and apparatus for generating light that can be used directly for photomedicine applications.
The foregoing and other problems are overcome and the object of the invention is realized by methods and apparatus in accordance with embodiments of this invention.
This invention teaches the use of static or flowing chemiluminescent materials to excite a photosensitive drug or to irradiate a tissue for performing photomedicine. Many reactions can be used for chemiluminescence, and include luciferase (Firefly reaction) and the oxalate ester/hydrogen peroxide (H2 O2) system. The latter reaction is commonly found in so-called "light sticks".
The fluorescers in the chemiluminescent reaction can be selected so as to emit exactly under the absorption band of many photosensitive drugs, such as those used in PDT, or those found in living systems, such as blood.
For example, rhodamine derivatives can be used for exciting the photoresponsive drugs Photofrin, ALA, PP IX, and FOSCAN, a photodynamic therapy related drug.
In order to sustain high luminous fluxes, the reaction can be created in a reservoir with a constant input of reactants and pumped through a catheter or a fluid containing externally applied patch, depending on the procedure. Catheters are ideally suited for treating, by example, esophageal cancer, while patches are ideally suited for treating skin conditions, as well as head and neck cancers. The reactants can be heated to increase the reaction rates and produce higher optical power.
The use of patches for skin and oral cavity treatments can eliminate the need for anesthesia, and furthermore can allow the patient to continue the treatment outside of the clinical setting. Patches may be made to operate in a similar manner to light sticks, and can be initiated by a pressure rupture of a barrier, releasing the reactants into a transparent pouch. One side of the pouch may be reflective so as to enhance the delivery of light towards the target area.
Active feedback can be provided from the target area using, by example, a photodiode, so as to control the delivery of reactants to, and the output power of the reactants at, the target area.
In preferred embodiments of this invention a plurality of reactants are mixed for forming a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths. The mixing of the reactants causes an excitation of the at least one selected fluorescer by energy transfer from an energetic key immediate in the chemiluminescent reaction to provide a fluorescent emission in the predetermined band of wavelengths.
The above set forth and other features of the invention are made more apparent in the ensuing Detailed Description of the Invention when read in conjunction with the attached Drawings, wherein:
FIG. 1 is block diagram of a system for delivering a flowing chemiluminescent source to a region of interest;
FIG. 2 illustrates a first embodiment of a patch suitable for being connected to the system of FIG. 1;
FIG. 3A illustrates a top view of a second embodiment of a patch, wherein reactants can be mixed on demand for generating the chemiluminescent source;
FIG. 3B illustrates the cross-sectional side view of the second embodiment of the patch in FIG. 3A, wherein reactants can be mixed on demand for generating the chemiluminescent source;
FIG. 4 illustrates an embodiment of a catheter that is useful for esophageal and other applications; and
FIG. 5 is a block diagram of the system shown in FIG. 1, wherein active feedback is used.
Referring to FIG. 1, there is illustrated a system 10 for flowing a chemiluminescent source. System 10 includes a reservoir 12 for containing a volume of reactants 14. Coupled to the reservoir 12 are a plurality of reactant sources 16a, 16b which provide, either continuously or on demand, measured amounts of reactants to the reservoir 12 so as to provide a desired intensity of emitted light. A heater 18 can be provided for heating the reactants 14 so as to increase the reaction rate, to produce a higher optical output, as well as to control the optical power. The reactants include at least one selected fluorescer, such as a rhodamine derivative, that is selected for its emission of a wavelength that optically excites a photosensitive drug of interest. The reactants also include materials known for use in providing chemiluminescence, such as luciferase and the oxalate ester/hydrogen peroxide (H2 O2) system.
Suitable fluorescers for use with this invention include the following:
Coumarin 440
9,10-diphenylanthacene (DPA)
Rhodamine 590
Rhodamine tetrafluoroborate
Sulforhodamine 640
Kiton red 620
Acridine orange
The enhancement capacity of TiO2 for different dyes (Ester, dye+Peroxide with and without TiO2 was confirmed for the following:
Rubrene (yellow emission)
9,10-bis(phenylethynyl)anthracene (yellow)
Coumarin 480 (blue)
In the foregoing examples, the number following the compound name indicates the emission wavelength.
The teaching of this invention is believed to produce satisfactory results using peroxyoxalate chemiluminescent systems (PO CL). In these systems, oxalic acid derivatives react with hydrogen peroxide in the presence of a fluorophore to produce a bright emission characteristic of the fluorescer. This reaction proceeds via an energetic key intermediate, which is proposed to be 1, 2-dioxetanedione.
Other oxalate esters include the following.
Bis(2-nitrophenyl)
Bis(4-nitro 3-trifluoromethyl)
Bis(4-nitro-2-formylphenyl)
Bis(4-nitro-2,6-dichlorophenyl)
Bis(2.4-dinitrophenyl)
Bis(2,5-dinitrophenyl)
Bis(2,4-dichlorophenyl)
Bis(pentacholorophenyl)
Bis(pentafluorophenyl)
Bis(3-trifluoro-methylphenyl)
Bis(3.5-di(trifluoro-methylphenyl)
Bis(2,6-dimethylphenyl)
Bis(4-methoxyphenyl)
Diphenyl
Phenylene
Bis(2-naphthyl)
Di-i-butyl
Bis(2-cyano-2-propyl)
Bis(2,2,2-trifluoro-ethyl
Bis(diphenylmethyl)
Another oxalate ester of interest is bis(2,4,6-trichlorophenyl) oxalate.
In an exemplary embodiment of this invention a Rhodamine derivative is excited by energy transfer from an energetic immediate in the chemiluminescent reaction, and provides a fluorescent emission in a desired band of wavelengths. A second emitter (e.g., a dye, chelate, or phosphor) can be used to coat a portion of structure, such as the catheter described below, to further shift the emission to a desired wavelength. A pump 20 is provided for pumping a flow of the reactants 14 through a conduit, such as a catheter 22, for delivering the chemiluminescent source to a region of tissue to be treated, as well as to control the output optical power. At least one portion of the catheter 22 is transparent to the light emission having the wavelengths used to excite a photosensitive drug of interest. FIG. 1 illustrates the use of a single transparent region 22a for outputting light A having the desired wavelengths.
It can be appreciated that the reactants can be pumped into a waste tank (not shown), and need not be returned to the reservoir 12.
FIG. 2 shows an embodiment wherein a patch 24 is inserted into the catheter 22 such that the flow of reactants passes through the patch 24. The patch has at least one internal chamber or compartment capable of containing the flow of reactants as they pass through. The use of the patch 24 beneficially spreads the flowing chemiluminescent source over a wider area, which is useful for treating certain tissues such as skin. As in the catheter embodiment of FIG. 1, the patch 24 has at least one region 24a that is transparent to the light A having the wavelengths of interest.
FIG. 3A illustrates a top view and FIG. 3B a cross-sectional side view of an embodiment wherein a partially transparent patch 26 includes a plurality of reactant compartments or containers 26a, 26b which, when broken or ruptured by a mechanical deformation of the patch, enable the reactants to be mixed and thus provide the desired therapeutic light. Although two reactants are shown in FIGS. 1 and 3A, it can be realized that as many reactants as are needed can be utilized in accordance with this invention. At least one of the reactants also contains the one or more selected fluorescers, or the fluorescer(s) could be provided in a separate compartment or container.
As is seen in FIG. 3B the patch 26 includes a transparent window portion 26c, and may include a reflector 26d, such as a layer of foil, disposed on an opposite surface for reflecting incident light towards the window portion 26c. The reflector 26d can be used as well in the embodiments of FIGS. 1 and 2. A portion of the reflector 26d, or some other structure in the patch 26, could include the above-mentioned second emitter (e.g., a dye, chelate, or phosphor) in order to shift the emission to a desired wavelength.
FIG. 4 illustrates an embodiment of a transparent or partially transparent catheter 30 that is useful for esophageal and other applications. A flow of reactants is established through the catheter 30 for providing the desired light emission A. A set of heating coils or tape 32, such as a resistive heating strip, can be used to produce a desired non-uniform excitation profile to treat different regions with dose selectivity. The coils 32 are preferably of low wattage, and may be molded into or embedded within the catheter wall. The reflector 26d can also be used if desired.
FIG. 5 shows the embodiment of FIG. 1 wherein a photodiode (PD) is used to provide an in-situ measure of output light dosage. The output of the photodiode is coupled to a controller 40 having an associated control bus 40a. The control bus 40a, in this example, is coupled to the pump 20, to reactant source pumps 16c and 16d, and to a heater controller 18a (shown schematically as a switch). Based on the light intensity detected by the photodiode in the target area, the controller is enabled to selectively control one or more reactant parameters (e.g., temperature, concentration, flow rate, etc.) so as to provide a desired amount of light energy to the target area. Other optical detectors, such as photoresistors, can be used in place of the photodiode.
While the invention has been particularly shown and described with respect to preferred embodiments thereof, it will be understood by those skilled in the art that changes in form and details may be made therein without departing from the scope and spirit of the invention.
1. An optical source for exciting a photosensitive drug that responds to light within a predetermined band of wavelengths, comprising a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths;
wherein said chemiluminescent source, in combination with said at least one fluorescer, is provided as a plurality of reactants, and further comprising:
a reservoir wherein said reactants are mixed; and
a pump for pumping the mixed reactants through a conduit to a tissue to be treated.
2. The optical source as in claim 1, wherein the chemiluminescent source is flowed during use.
3. The optical source as in claim 1, and further comprising means for heating said chemiluminescent source.
4. The optical source as in claim 1, and further comprising an active feedback system for controlling said output light.
5. The optical source as in claim 1, wherein said chemiluminescent source in combination with said at least one fluorescer are flowed through a conduit.
6. The optical source as in claim 1, wherein said chemiluminescent source is comprised of an oxalate ester/hydrogen peroxide (H2 O2) system.
7. The optical source as in claim 1, wherein said chemiluminescent source is comprised of luciferase.
8. The optical source as in claim 1, wherein said at least one selected fluorescer is comprised of a rhodamine derivative.
9. The optical source as in claim 1, and further comprising a second fluorescer selected for shifting emission wavelengths of said at least one fluorescer.
10. An optical source for exciting a photosensitive drug that responds to light within a predetermined band of wavelengths, comprising a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths;
a patch comprising a plurality of compartments, individual ones of said compartments containing one of said reactants; wherein
a mechanical deformation of said patch causes said compartments to rupture such that said reactants are released and mixed together.
11. The optical source as in claim 10, wherein said chemiluminescent source is comprised of at least one of an oxalate ester/hydrogen peroxide (H2 O2) system or of luciferase.
12. The optical source as in claim 10, and further comprising a second fluorescer selected for shifting emission wavelengths of said at least one fluorescer.
13. A catheter for insertion into a body cavity, said catheter having at least one region that is transparent to light having wavelengths within a predetermined band of wavelengths selected for exciting a photosensitive drug, said catheter comprising a conduit for flowing a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths.
14. A patch for application to a surface of a tissue, said patch having at least one region that is transparent to light having wavelengths within a predetermined band of wavelengths selected for exciting a photosensitive drug, said patch comprising an internal chamber region for flowing therethrough a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths.
15. A patch for application to a surface of a tissue, said patch having at least one region that is transparent to light having wavelengths within a predetermined band of wavelengths selected for exciting a photosensitive drug, said patch having at least one region that is transparent to light having wavelengths within a predetermined band of wavelengths selected for exciting a photosensitive drug, said patch comprising a plurality of compartments, individual ones of said compartments containing one of a plurality of reactants; wherein a mechanical deformation of said patch causes said compartments to rupture such that said reactants are released and mixed together for forming a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths.
16. A method for generating light having wavelengths within a predetermined band of wavelengths selected for exciting a photosensitive drug, comprising steps of:
mixing a plurality of reactants for forming a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths;
exciting the at least one selected fluorescer by energy transfer from an energetic immediate in a chemiluminescent reaction to provide a fluorescent emission in the predetermined band of wavelengths; and
optically coupling the fluorescent emission to a tissue to be treated by the photosensitive drug.
17. The method as in claim 16, wherein the step of mixing includes a step of mixing the reactants in a reservoir, and wherein the step of optically coupling includes a step of causing a flow of the chemiluminescent source in combination with the at least one fluorescer to the tissue to be treated.
18. The method as in claim 16, wherein the step of mixing includes a step of combining the reactants within a patch that is to be applied to a surface of a tissue to be treated.
19. The method as in claim 18, wherein the step of combining includes an initial step of rupturing a plurality of reactant containing compartments within the patch.
20. A source of light having wavelengths within a predetermined band of wavelengths selected for exciting a photosensitive drug, comprising:
means for mixing a plurality of reactants for forming a chemiluminescent source in combination with at least one fluorescer selected to output light within the predetermined band of wavelengths, said mixing means causing an excitation of the at least one selected fluorescer by energy transfer from an energetic immediate in a chemiluminescent reaction to provide a fluorescent emission in the predetermined band of wavelengths; and
means for optically coupling the fluorescent emission to a tissue to be treated by the photosensitive drug.
21. The source as in claim 20, wherein said mixing means is comprised of:
a reservoir coupled to a plurality of reactant sources;
means for introducing reactants from said plurality of reactant sources into said reservoir; and
means for causing a flow of mixed reactants out of said reservoir.
22. The source as in claim 20, wherein said means for optically coupling is comprised of a catheter having a transparent region.
23. The source as in claim 20, wherein said means for optically coupling is comprised of a patch having a transparent region.
24. The source as in claim 23, wherein said mixing occurs within said patch.
25. The source as in claim 20, and further comprising means for maintaining said chemiluminescent source at a predetermined temperature.
26. The source as in claim 20, and further comprising an active feedback system for controlling an intensity of said fluorescent emission.
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Owner name: SPECTRA SCIENCE CORPORATION, RHODE ISLAND
Free format text: ASSIGNMENT OF ASSIGNORS INTEREST;ASSIGNOR:LAWANDY, NABIL M.;REEL/FRAME:008815/0010
2006-12-08 REIN Reinstatement after maintenance fee payment confirmed
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text:
The Problem: Violence against women
The Treaty: Global Game Changer
How a Treaty is Born
Treaty Development
Violence Against Women and the Law
The People's Call
Francisco Rivera, JD, LLM, Puerto Rico/USA
Steering Committee Member
Francisco Rivera, JD, LLM, Puerto Rico/USA, is a human rights attorney, law professor, and founding director of the International Human Rights Clinic at Santa Clara University in California. He specializes in the Inter-American Human Rights System and was a senior staff attorney at the Inter-American Court of Human Rights of the Organization of American States, seated in Costa Rica. He has been a consultant for a number of nongovernmental organizations (NGOs), as well as for the United Nations Development Programme (UNDP), the International Labor Organization (ILO), and the Inter-American Institute of Human Rights (IIHR). In the late 1990s, he served as executive director of the Amnesty International Section in Puerto Rico. Francisco joined the Every Woman Treaty Working Group in 2015 and chaired the Expert Special Committee on Governing Bodies.
"Yo firmé porque un nuevo tratado generará el momentum necesario para obtener nuevos recursos, ideas, energías, y voluntad política para combatir la violencia contra la mujer."
"I signed because a new treaty will generate the momentum necessary to obtain new and additional resources, ideas, energy, and political will to eradicate violence against women."
- Francisco Rivera
Next member >
The next step in ending violence against women and girls.
Every Woman Treaty is a 501(c)(3) organization | EIN/tax ID: 47-3272024


On those occasions that I head out with purely the intent to browse or return items, instantly evolve into the days that I end up buying stuff. Good buys, but unintended, none the less.
But, trust me, if I head out for the purpose of buying some new pieces and looking for something special, that is GUARANTEED to be the day that I spend hours roaming aimless and bagless.
I happened upon this skirt on one of those non-shopping occasions. I had returned some items to one store and just stopped off in Forever 21 to take a looksy. And there was this darling skirt in on a rack by its lonesome in a section in which it did not even belong.
My size and $20. Sold!
The salesgirl even told me this was the only one they'd received and us fashion girls know how special that is!
It is simply one of the cutest pieces I have in my wardrobe and I've held on to it for some time waiting for just the right kind of top to pair with it. That "perfect" top never came and I ran out of patience, so this time I wore a dress as a shirt for a more casual feel. Re-purposing items in your closet to do double duty is always awesome, right?!


text:
Apple Said To Be Developing Contact-Free Wireless Charging for iPhone
Filed to: appleFiled to: apple
Apple apparently wants to help you charge your iPhone no matter where you are in your home. A new report suggests that it's building a new wireless charging technology that wouldn't require direct contact with a power mat.
Bloomberg reports that Apple "is exploring cutting-edge technologies that would allow iPhones and iPads to be powered from further away than the charging mats used with current smartphones."
The idea is not a new one. Earlier this month, Ossia showed off a system that allows an iPhone to be charged in mid-air. It works by using an antenna that can capture energy held in radio frequency waves floating through the air that are carefully directed from a central hub.
It's not an easy challenge, for a number of reasons. The technology required to capture enough energy is, for Ossia at least, rather bulky. And as Bloomberg points out, "efficiency of power transfer decreases as the distance between transmitter and receiver grows." If Apple is trying to build a sleek phone, those two issues create a compounded problem where it's difficult to charge a battery quickly.
Indeed, Bloomberg claims that Apple "is looking to overcome technical barriers." No kidding. In fact, it's likely been working on wireless charging for years by this points. Whether we'll see in the next iPhone, or the one after that? We'll have to wait and see.
[Bloomberg]
Photo by Michael Hession
Recent from Jamie Condliffe
Engineers Own the Future—and Maybe Even Us
Tiny Metallic Pixels Could Make Pictures That Never Fade
The Beautiful Birth of a New Star


text:
SMark Sailing
SMark Swimming
Gilles Caprari
With a PhD in robotics at the Eidgenössische Technische Hochschule Zürich ETH, an entrepreneurial career and years of sailing experience, Gilles is the tech genius behind SMark.
It doesn't take long for him to bring to life anything you might ask for which did not exist before.
His precious expertise brings WaterSportMarks at the forefront of water drones. In 2020, after working with Julien and Valentina, Gilles founded WaterSportMarks together with them to work on multiple versions of robotic marks.
Valentina Venturi
During a long mark resetting in a regatta in 2016, Valentina started to dream of a light drone mark that even a girl could could handle alone.
After a few years building prototypes she met Gilles and Julien in 2020, they were working as well on a light drone buoy... finally the dream became reality.
With a PhD in mechanical engineering and the past 8 years spent working at the European Center for Nuclear Research Valentina is now fully focussed on robotic marks!
Julien Buros
Either it's behind a boat, under a kite or on a snow slope, Julien is a 'water' sport fan! He is a waterski instructor and holds a Msc. in electrical engineering from Polytechnic Grenoble (FR) and University of Florida. Julien has been designing GPS systems for almost 10 years and brings 20 years of experience in high-tech companies ranging from start-up to multinationals. SMark was first born in his mind as a waterski robotic mark but Gilles and Valentina's sailing influence got the upper hand…for now!


text:
Fan Tai (355– 21 September 428), courtesy name Bolun, was an official who served under the Eastern Jin and Liu Song dynasties of China. His fourth son, Fan Ye, wrote the historical text Book of the Later Han.

References

 The Cambridge History of China, Volume 1, The Ch'in and Han Empires, 221 BC–AD 220 https://web.archive.org/web/20070927223220/http://www.cnread.net/cnread1/lszl/s/shenyue/ss/060.htm (Chinese only)

355 births
428 deaths
Jin dynasty (266–420) generals
Liu Song generals
Chinese educators
Jin dynasty (266–420) politicians
Liu Song politicians
Jin dynasty (266–420) essayists
Northern and Southern dynasties essayists
Liu Song writers

redpajama_set_name: RedPajamaWikipedia