|
1.IntroductionThe genus Fusarium is a well-known plant pathogen that causes damage to agriculture.1,2 Over the past 30 years, the Fusarium species have been intensively studied and of the 61 identified species, at least 35 are known to produce mycotoxins that are harmful to animals and humans.3–8 The Fusarium mycotoxins are known as chemically different secondary metabolites. These mycotoxins can contaminate cereal grains causing disease and death in the animals that consume them. The trichothecenes [T2 toxin, deoxynivalenol, nivalenol, fusarenon-X (FX), and diacetoxyscirpenol (DAS)], fumonisins, zearalenone, and moniliformin are the most important Fusarium mycotoxins.9,10 These mycotoxins cause neurological damage and are frequently associated with animal and human diseases.4,5 Several mycotoxins are stable under normal food-processing conditions; thus, they may be present in natural foods and processed products.11,12 Individuals who consume contaminated food are not exposed to a single mycotoxin; rather, they are exposed to a cocktail of them.13 In vitro experiments have shown that a mixture of mycotoxins results in additive toxicity or synergy that reduces the effective toxic dose of each component in the mixture.14,15 If this potentiating occurs in vivo, then the health consequences will be more severe. In humans, the species Fusarium may cause local, invasive, or disseminated infections. The pathogen typically affects immunocompromised individuals because infections in immunocompetent individuals are reported less frequently.16,17 Among immunocompromised patients, Fusarium is the second most common cause of fungal infection, with an increased incidence in patients with hematologic malignancies and especially in those exposed to recipients of hematopoietic stem cell transplantations.18 The entry points to the disseminated infection include the respiratory tract, gastrointestinal tract, and skin.16 Skin lesions, which are typically located on the extremities, are often described as painful subcutaneous nodules that may ulcerate and become necrotic lesions of the gangrenous erythema type.18 For skin infections, the entry points are wounds, digital ulcers, onychomycosis, and paronychia.17,19–21 In recent years, the number of Fusarium onychomycosis cases in immunocompetent patients has increased significantly.17,22 Guilhermetti et al.17 reported a 7.5% incidence of confirmed cases of onychomycosis by Fusarium spp. and Fusarium oxysporum (F. oxysporum). The latter is one of the most common and pathogenic of the genus Fusarium and is the specie that has been mostly isolated.18,22 In humans, this specie is an important emerging pathogen due to the increasing number of severe cases reported and its wide-ranging resistance to antifungal drugs.23,24 The increase in F. oxysporum infections in immunocompetent patients is an important aspect; thus, the evaluation of the biological and physicochemical effects of F. oxysporum on the skin of healthy rats and their relation with induced apoptosis is of utmost importance. It is well recognized that techniques based on infrared radiation provide an important route to access chemical bonding information of biological tissues.25–30 This can be done by exploring the finger print characteristics of the optical absorption bands in this spectral region. Among these methods, Fourier transform infrared photoacoustic spectroscopy (FTIR-PAS) has the special characteristic of allowing depth profile studies to detect the penetration and interaction of substances in biological tissues.31–33 It can be applied for measurements at clinical conditions because minimal sample preparation is needed. It is a nondestructive method and permits inspection in opaque and highly scattering samples. Therefore, the aim of this study was to investigate the biological and physicochemical effects of a crude extract (CE) of F. oxysporum on the skin of healthy rats. The FTIR-PAS spectroscopy was applied to evaluate the induced physicochemical structural changes and their relation with the occurrence of apoptosis. 2.Materials and MethodsThe study protocol was reviewed and approved by the Local Ethics Committee for Animal Experimentation of the Universidade Estadual de Maringá (UEM), under protocol number 005/2010 and statement 080/2010. 2.1.Fungal Isolate and Culture ConditionsThe F. oxysporum sample used in this study was previously isolated from an onychomycosis patient at the Teaching and Research Laboratory of the Departamento de Ciências Básicas da Saúde, UEM, Maringá, Paraná, Brazil. The fungus was peaked in Petri dishes containing potato dextrose agar and incubated for 7 days at 25°C. Three 5-mm discs were removed from the culture and placed into flasks containing 200 mL of Czapek–Dox (CZ) medium, which had been adjusted to a pH of 5.5 and sterilized at 120°C for 20 min. The flasks were maintained at a 70-rpm orbital agitation during 15 days at 25°C. Next, the culture was sterilized at 120°C for 20 min and filtered with Whatman filter paper No. 1 for mycelium removal. The filtration product was sterilized on a Millipore filter with a 0.45-μm pore size and dialyzed against the distilled water overnight. The obtained extract, referred to as CE, was maintained at 4°C until use. As an experimental control, 100 mL of CZ medium were produced and exposed to the same condition as that used for the fungus-containing medium. 2.2.Experimental ProceduresMale Wistar rats weighing 150 to 200 g were used. The animals were kept in polypropylene boxes at a ratio of four animals/box in an environment with a controlled temperature of , 50% humidity, and a light-dark cycle of 12 h, with water and food ad libitum. While under anesthesia with sodium thiopental (), an area of on the back, near the cervical region of each animal, was epilated 2 days before the topical application of the solutions. This procedure was adopted in order to avoid inflammatory reaction that would interfere in the results interpretation. On the day of the experiment, 50 μL of the CE or 50 μL of the CZ medium were topically applied to each animal of the experimental and control groups, respectively. The animals were sacrificed 3, 6, 12, and 24 h after the application of the solutions. 2.3.Physicochemical Analysis of the Skin by FTIR-PASFor the physicochemical analysis of the skin following topical application of either the CE or the CZ medium, two animals/time were used for each experimental group. After each animal sacrifice, the skin containing both epidermis and dermis was removed and sent immediately for ex vivo analysis using an FTIR spectrometer (Varian, Model 7000) equipped with an MTEC model 300 photoacoustic detection cell. Each skin sample was positioned inside the cell chamber with its epidermal side upside down allowing the radiation to excite the dermal region, the opposite side in relation to where the CE or the CZ medium was applied, as illustrated in Fig. 1. After that the cell chamber was purged with the helium gas for approximately 5 min to remove air moisture, and subsequently it was sealed to perform the measurements. This procedure increases the sensitivity of the technique and minimizes spectral interference from loosely bonded molecules of the air such as . All the spectra were collected in the rapid scan mode in the spectral range between 4000 and , with resolution, an average of 28 scans and the interferometer mirror velocity () of (10 kHz). The reference spectrum was acquired by the use of a carbon black sample. The depth analysis in the sample was estimated through the thermal diffusion length (cm) as , in which is the sample thermal diffusivity () and is the incident radiation wavelength (cm). This parameter is the dimension over which the thermal wave decays to of its original amplitude and can be used in the analysis as approximately the sampling depth where the PAS signal is generated. In our experimental conditions, using the skin thermal diffusivity measured before as (Ref. 34) and , we can calculate the values of μ according to the incident radiation wavelength, being, e.g., 2.8 μm at and 1.7 μm at . Therefore, since the exciting radiation was incident on the dermal side of the skin and the application of the CE and CZ medium was done on the epidermal side, the detection of optical bands would be an indication that the formulation penetrated throughout the skin. 2.4.Histological StudyThe histological studies were performed in four animals for each time interval after the formulations’ application in both experimental and control groups. After killing, the skin was collected and fixed in 4% paraformaldehyde for a period of 24 h. The skin was then processed for paraffin embedding. Semi-serial 7-μm thick sections were prepared and placed on slides. The samples were used for the following techniques: (a) hematoxylin and eosin (H&E) staining for the morphological observation and morphometric analysis of the epidermis and dermis thickness; (b) sirius red (picro sirius) staining for the analysis of the area occupied by collagen in the dermis; (c) immunohistochemical in situ detection of fragmented DNA [terminal dUTP nick end labeling (TUNEL assay)] for the analysis of apoptosis in the epidermis and dermis; and (d) immunohistochemistry using anti-proliferating cell nuclear antigen (PCNA) to study the cell proliferation in the epidermis and dermis. 2.4.1.Histomorphometric measurementsThe skin morphology was observed microscopically to study the occurrence of histopathological alterations, such as the presence of inflammatory infiltrations or epidermis or dermis structure alterations. The inflammatory reaction was graded as absent (−), weak (+), moderate (+ +), or severe (+ + +) in accordance with the frequency of cells in the dermis. Ten histological sections/animals were analyzed. The observations were performed with an Olympus BX41 microscope adopting the double-blind method. The two different researchers who made the observations did not know which slides contained the control or the treated skins. For the morphometric analysis of the epidermis and dermis thicknesses, histological section images were captured using an Olympus BX41 optical microscope with a objective coupled to a QColor 3 Olympus camera. The measurements were acquired using the Image Pro Plus® software, version 4.5 (Media Cybernetics). The images from 10 sections/animals were analyzed, and 200 measurements/animals were separately recorded for the epidermis and dermis. The results are expressed as the . 2.4.2.Analysis of area covered by the collagen fibers of the dermisThe analysis of the collagen fibers of the dermis was performed by light microscopy using optical polarization. This method based on birefringence allows the collagen to be classified as type I or III. For the morphometric analysis of the area covered by collagen, standardized images of 12 fields of were captured with a objective. The image capture was performed with the same illumination intensity and was digitized and analyzed with the Image Pro Plus® software (Version 4.5, Media Cybernetics). The results are expressed as the in . 2.4.3.In situ detection of fragmented DNA—terminal dUTP nick end-labeling (TUNEL assay)DNA fragmentation was examined using an apoptosis detection kit (ApopTag® Peroxidase, Chemicon). After deparafinization and rehydration, the slides containing the histological sections were subjected to enzymatic digestion with of proteinase K for 5 min, treated with 5% hydrogen peroxide (30 volumes) for 20 min to block endogenous peroxidase, and washed with 0.1 M PBS with pH 7.4. The sections were then immersed in equilibration buffer for 10 min and incubated with a solution containing the enzyme terminal deoxynucleotidyl transferase at 37°C for 1 h. The sections were washed in PBS and incubated with the conjugated anti-digoxigenin peroxidase and peroxidase substrate to detect apoptosis signs stained in brown. Counterstaining was performed with Harris hematoxylin. The immunostaining on the epidermis and dermis was performed based on the frequency of the cells stained adopting the following classification: absent (−), weak (+), moderate (+ +), or strong (+ + +). 2.4.4.Identification of cells under proliferation (PCNA)The detection of cells undergoing proliferation was performed using the primary antibody anti-PCNA (Zymed). After deparafinization and rehydration, the histological sections were subjected to antigenic recovery in a citrate buffer with a pH of 6.0. The primary antibody anti-PCNA () was incubated for 60 min at room temperature (RT). A commercial kit (Histostain® Plus Mouse Primary DAB Zymed) was used at RT for detection. The sections were counterstained with Harris hematoxylin. The immunostaining on the epidermis and dermis was classified as absent (−), weak (+), moderate (+ +), or strong (+ + +) based on the frequency of the cells stained. 2.4.5.Statistical analysisThe results of the collagen-covered area and skin thicknesses were subjected to statistical treatment. All the data were analyzed with Kolmogorov–Smirnov, D’Agostino and Pearson, and Shapiro–Wilk normality tests. When the data did not have a normal distribution, an analysis of variance and the Kruskall–Wallis nonparametric test along with the Dunn post test were used. A 5% significance level was used (). 3.Results and DiscussionThe obtained CE is a complex mixture produced by fungus growth, which includes metabolism products such as secondary metabolites like mycotoxins. The extract also contains partially digested culture medium residue components. The spectral analyses of organic compounds by FTIR represent a method for obtaining a “fingerprint” of the molecules, i.e., allowing for their identification from the recognition of the functional groups to which they belong. The bands between 1700 and have been related to amides I and II, and they are frequently used as an indicator of secondary protein structure.35,36 The bands between 3000 and are related to the symmetric and asymmetric stretching of groups in proteins and lipids36,37 and those between 3800 and to the and amine () molecules.36 It is known that the identification of amides I and II and groups may allow the monitoring of conformational changes detected by the vibrational variations of these molecules when subjected to a state different from that considered in physiological tests.38 Figures 2(a) and 2(b) show the FTIR-PAS absorption spectra between 1900 and of the dermis of rats without the application of any substance and at 3, 6, 12, and 24 h after the topical application of both CZ culture medium or F. oxysporum CE. The changes in the absorption bands suggest structural alterations of amide I (1720 to ), amide II (1580 to ) and (1470 to ) groups, which occurred in the dermis after the topical application of the F. oxysporum CE. These spectral modifications may imply that the dermis underwent physicochemical changes that were not observed in the untreated normal skin or the skin on which the culture medium was applied. The region around 1900 to was more intense for the 3 and 24 h control samples. One possible explanation for this difference may be the presence of higher amounts of adipose tissues in the illuminated areas. We should stress that this spectral region was not considered in our interpretation. Moreover, Fig. 3 shows the FTIR data in the spectral region between 3850 and . It is observed that the and amine () bands presented an increase in their intensities with time after the application of both CE and CZ medium. Once these bands are present in both, this result confirms that the CE and CZ medium permeate through the skin reaching the dermal region. Therefore, our hypothesis is that the physicochemical changes in the skin dermal side observed from the FTIR-PAS data strongly suggest that either partial or complete extract permeation occurred and then produced the observed skin structural alterations. Figure 4 shows the behavior of the peak area at approximately within the group region (1350 to ) for 3, 6, 12, and 24 h after the topical application of the F. oxysporum CE. The control point () represents the peak area of the dermis without the application of any substance. In the infrared region, the group also gives rise to absorption in the region. At this frequency, the molecule has an asymmetric bending vibration mode. This group also has vibrational characteristics at , which is a symmetric bending vibration mode.36 The spectra obtained for the epidermis and dermis after the application of the CZ medium showed different behavior from that observed after the application of the CE, suggesting that the medium and extract have different contributions to the potential structural changes of the highlighted groups (amide I, amide II, and ) as evidenced in Fig. 2(b). Histopathological evaluation did not show macroscopic changes on the skin surfaces of the rats treated with the F. oxysporum extract. However, during the microscopic examination of the sections stained with H&E, the following changes were detected: (1) red cells were observed on the surface of the skin (at 3 and 6 h), (2) keratinocytes were present on the epidermis with a bright, perinuclear halo that is typical of hydropic alterations, mainly at 3 h, and (3) the cell number increased on the dermis, and the cells were diffusely localized, particularly for the higher frequencies of fibroblasts and macrophages, but few neutrophils and no formation of typical inflammatory infiltrates were observed, as shown in Fig. 5. The inflammation score is shown in Table 1. Table 1Score representing the frequency of skin inflammatory cells 3, 6, 12, and 24 hs after the topical application of the Czapek–Dox (CZ) medium or the F. oxysporum crude extract (CE).
(−) absent, (+) weak, (+ +) moderate, and (+ + +) severe. Skin morphometry analysis showed the epidermis of the animals treated with the CE were thicker () after 6, 12, and 24 h [Fig. 6(a)]. The dermis was thicker after 12 and 24 h [Fig. 6(b)]. For the animals treated with the CE and death after 3 and 6 h, the thicknesses of the dermis were significantly reduced [Fig. 6(b)]. Bhavanishankar et al.39 suggested that several toxins produced by Fusarium act in different manners when they appear in different combinations. The authors evaluated the variations in epidermis thickness in vivo in guinea pigs after the topical application of isolated toxins or a combination of toxins produced by F. oxysporum. The epidermal thickening by T2 toxin combined with either FXor butenolide (Bd) was greater than by T2 alone. This finding indicates an antagonistic relationship between these toxins. A greater thickness occurred upon the application of a combination of DAS, which is a trichothecene toxin, and Bd, which is a nontrichothecene toxin, suggesting a synergistic toxic effect by this combination. In the dermis of the animals treated with the CE, the area covered by collagen type I increased after 3, 6, and 12 h, and the area covered by collagen type III increased after 3, 6, 12, and 24 h compared to the dermis of the animals treated with the culture medium [Fig. 7(a)]. For all the evaluated time points, the total area covered by collagen was larger () in the groups treated with the CE [Fig. 7(b)]. Figure 7(c) shows the proportion of collagen III:I. The greatest disparity in the ratio of the collagens fibers (6 and 24 h) occurs after the maximum for death (3 h) and proliferation (12 h) of fibroblasts. With the death of cells (fibroblasts), the matrix is reabsorbed. A slight decrease in the mature collagen, type I, is observed in skin treated with the CE. With the formation of new cells, a matrix is produced, and the increase in the immature collagen, type III, is observed. In the control group, a balance in the proportions of the types I and III fibers could be observed. TUNEL immunohistochemical reaction to the stained apoptotic cells results are shown in Table 2 and Figs. 8(a) and 8(b). The basal layer of the epidermis showed the highest frequency and positive intensity after labeling with TUNEL, followed by the Prickle-cell layer and the other layers that had more diffuse labeling. In the dermis, fibroblasts and hair follicles cells were the most immunostained cell types. The strong staining of the follicles was most likely due to its involutive process that occurred after epilation.40 A large number of stained cells were observed in the skin tissue and panniculus carnosus muscle cells. In a previous study, we reported the occurrence of programmed cell death in these same tissue types after the intradermal administration of a F. oxysporum CE that was grown under the same conditions.41 These results demonstrated the potential permeation capacity through the skin and the harmful effects of this extract, even after a single application. Table 2Terminal dUTP nick end labeling immunostaining score of the epidermis and dermis 3, 6, 12, and 24 h after the topical application of the Fusarium oxysporum CE or CZ culture medium on the skin of Wistar rats.
(−) absent labeling, (+) weak labeling, (+ +) moderate labeling, and (+ + +) strong labeling based on the frequency of the stained cells. The capacity of apoptosis induction on basal cells within the epidermis after a topical application of the Fusarium T2 toxin was shown in a model of Wistar rats WBN/ILA-Ht.42 Albarenque and Doi43 found a 40% decrease in the viability of keratinocytes in primary culture after 3 h, with evolution up to 12 h after the administration of T2 toxin. These authors used an in vitro model. Morphological alterations (e.g., chromatin condensation and margination and nuclear fragmentation, all of which are characteristics of cells undergoing apoptosis) were also recorded by optical and electron microscopy. In particular, the potential of programmed cell death induction has been attributed to trichothecenes,42–46 which are molecules that have molecular weights of 250 to 500 Da.47 In the present study, using a dialysis membrane with a molecular weight cut-off of 12 to 16 kDa may have removed the trichothecenes from the CE; thus, the induction of apoptosis could be attributed to other extracted components with toxic potential. Another hypothesis includes the formation of complexes between smaller molecules that can reach molecular weights . These complexes would not be eliminated in the dialysis process and could maintain their toxic potential, inducing the apoptosis observed by TUNEL. It is important to observe that the period with higher intensity of TUNEL, 3 h after CE application, coincides with the most pronounced structural changes observed in the FTIR-PAS data, revealed by the modifications in the optical absorption bands associated with the group, at . A similar result was detected in the controls, suggesting that there were structural changes that could be related to the process of programmed cell death in the skin treated with the fungal extract. Marder et al.48 reported that the optical absorption band at was related to the presence of mycotoxins produced by B. sorokiniana. Table 3 and Figs. 8(c) and 8(d) show the results obtained after the immunohistochemistry reaction to detect anti-PCNA-positive cells. The results showed that there is a thickening of the epidermis within 6, 12, and 24 h after the application of the CE. The strong labeling of PCNA, the expression of which increased in the G1 and S phases of the cell cycle,49–52 indicates an increase in the proliferative activity of this tissue after 3 h and moderate labeling after 6, 12, and 24 h. However, as shown above, strong TUNEL was observed at 3 h, and a moderate level of labeling was observed after 6, 12, and 24 h. We believe that the epidermis thickening would reflect the changes in the kinetic activity of the keratinocytes, i.e., stimulating the proliferative activity because of an increased cell death after the topical application of the extract. As shown before, the skin is one of the entrance of Fusarium.16 It is able to break the epithelial barrier inducing cutaneous lesions known as fusariosis.18 Then, it is reasonable to suppose that the topical application of the CE may be able to temporarily modify the epithelial permeability resulting in cutaneous dehydration because of the higher concentration gradient of the CE in the dermis, with consequent changes in its thickness, as occurred 3 and 6 h after the extract application. In the same period, a number of keratinocytes showed hydropic changes, a reversible state resulting from cellular injury.53 Table 3Proliferating cell nuclear antigen immunostaining scores for the epidermis and dermis 3, 6, 12, and 24 h after the topical application of the F. oxysporum CE or CZ culture medium on the skin of Wistar rats.
(−) absent labeling, (+) weak labeling, (+ +) moderate labeling, and (+ + +) strong labeling. The skin response after topical application differed from that observed after the intradermal inoculation of the F. oxysporum CE. Although the number of inflammatory cells in the dermis increased, there was no formation of typical inflammatory infiltrate. The most frequent inflammatory cells were macrophages, followed by a small number of lymphocytes and neutrophils. These characteristics suggest that the epidermis has exercised its barrier function, preventing the extract from fully permeating. The dermal fibroblasts were largely responsible for the increased cellularity after the application of the Fusarium extract. Fibroblasts are the cells responsible for the production of extracellular matrix components. In vitro studies showed that these cells are susceptible to the action of mycotoxins.53,54 We observed that the dermal fibroblasts showed strong labeling for cell death after 3 h and moderate labeling in other periods. In contrast, the fibroblasts showed moderate positivity for proliferation at 3 and 6 h, a peak at 12 h, and decay at 24 h. Thus, these results indicate the pursuit of tissue component replacement after the imbalance caused by an external agent. The morphometric analysis of the dermis thickness and the area covered by collagen support this hypothesis because after 12 h, there was significant dermal thickening, which was most likely due to increased collagen synthesis [Fig. 6(b)]. 4.ConclusionIn this work, we demonstrated that the F. oxysporum CE could permeate the epidermis and dermis, reaching the subcutaneous tissue, inducing apoptosis of the cells in these tissues, and causing physicochemical changes in the organic molecules located in the dermis. Considering that the animals were immunocompetent, the proliferative and morphological changes observed in this study were derived from a process of tissue recovery after the irritating stimulation ended. We believe that the procedure used in this work may be useful for future studies in immunocompromised knockout animals to investigate the dynamics of the physicochemical changes that may occur in the infected tissues. AcknowledgmentsWe are thankful to Brazilian agencies CAPES, CNPq, FINEP, and Fundação Araucária for the financial support of this work. ReferencesJ. T. Mills,
“Ecology of mycotoxigenic Fusarium species on cereal seeds,”
J. Food Protect., 52
(10), 737
–742
(1989). JFPRDR 0362-028X Google Scholar
J. W. Rippon, Tratado de Micologia Médica, 3rd ed.Mc Graw Hill, México
(1990). Google Scholar
J. GuarroJ. Gene,
“Opportunistic fusarial infections in humans,”
Eur. J. Clin. Microbiol. Infect. Dis., 14
(9), 741
–754
(1995). http://dx.doi.org/10.1007/BF01690988 EJCDEU 0934-9723 Google Scholar
E. Conkováet al.,
“Fusarial toxins and their role in animal diseases,”
Vet. J., 165
(3), 214
–220
(2003). http://dx.doi.org/10.1016/S1090-0233(02)00127-2 VTJRFP 1090-0233 Google Scholar
J. Evanset al.,
“Intracranial fusariosis: a novel cause of fungal meningoencephalitis in a dog,”
Vet. Pathol., 41
(5), 510
–514
(2004). http://dx.doi.org/10.1354/vp.41-5-510 VTPHAK 0300-9858 Google Scholar
S. M. Abdel-Rahmanet al.,
“Pharmacokinetics of terbinafine in young children treated for tinea capitis,”
Pediatr. Infect. Dis. J., 24
(10), 886
–891
(2005). http://dx.doi.org/10.1097/01.inf.0000180577.29853.a0 PIDJEV 0891-3668 Google Scholar
M. L. D. F. PerónJ. J. V. TeixeiraT. I. E. Svidzinski,
“Epidemiologia e etiologia das dermatomicoses superficiais e cutâneas na Região de Paranavaí-Paraná, Brasil,”
Rev. Bras. Anal. Clin., 37
(2), 77
–81
(2005). RBACBX Google Scholar
J. A. A. Oliveiraet al.,
“Superficial mycoses in the City of Manaus/AM between March and November/2003,”
An. Bras. Dermatol., 81
(3), 238
–243
(2006). http://dx.doi.org/10.1590/S0365-05962006000300005 ABDEB3 0365-0596 Google Scholar
J. P. F. D’MelloC. M. PlacintaA. M. C. MacDonald,
“Fusarium micotoxins: a review of global implications for animal health, welfare and productivity,”
Anim. Feed Sci. Technol., 80
(3), 183
–205
(1999). http://dx.doi.org/10.1016/S0377-8401(99)00059-0 AFSTDH 0377-8401 Google Scholar
C. M. PlacintaJ. P. F. D’MelloA. M. C. MacDonald,
“A review of worldwide contamination of cereal grainsand animal feed with Fusarium mycotoxins,”
Anim. Feed Sci. Technol., 78
(1), 21
–37
(1999). http://dx.doi.org/10.1016/S0377-8401(98)00278-8 AFSTDH 0377-8401 Google Scholar
E. C. HopmansP. A. Murphy,
“Fumonisins: mycotoxins produced by Fusarium moniliforme,”
Natural Protectants Against Natural Toxicants, 61
–78 Technomic Publishing, Lancaster, Pennsylvania
(1993). Google Scholar
D. R. LaurenW. A. Smith,
“Stability of 328 Fusarium mycotoxins nivalenol, deoxynivalenol and zearalenone in ground maize under typical cooking conditions,”
Food Addit. Contam., 18
(11), 1011
–1016
(2001). http://dx.doi.org/10.1080/02652030110052283 FACOEB 1464-5122 Google Scholar
M. MarescaJ. Fantini,
“Some food-associated mycotoxins as potential risk factors in human predisposed to chronic intestinal inflammatory diseases,”
Toxicon, 56
(3), 282
–294
(2010). http://dx.doi.org/10.1016/j.toxicon.2010.04.016 TOXIA6 0041-0101 Google Scholar
E. E. Creppyet al.,
“Synergistic effects of fumonisin B1 and ochratoxin A: are in vitro cytotoxicity data predictive of in vivo acute toxicity?,”
Toxicology, 201
(1–3), 115
–123
(2004). http://dx.doi.org/10.1016/j.tox.2004.04.008 TXCYAC 0300-483X Google Scholar
J. H. Kouadioet al.,
“Effects of combinations of Fusarium mycotoxins on the inhibition of macromolecular synthesis, malondialdehyde levels, DNA methylation and fragmentation, and viability in Caco-2 cells,”
Toxicon, 49
(3), 306
–317
(2007). http://dx.doi.org/10.1016/j.toxicon.2006.09.029 TOXIA6 0041-0101 Google Scholar
A. K. GuptaR. BaranR. C. Summerbell,
“Fusarium infection of the skin,”
Curr. Opin. Infect. Dis., 13
(2), 121
–128
(2000). http://dx.doi.org/10.1097/00001432-200004000-00005 COIDE5 0951-7375 Google Scholar
E. Guilhermettiet al.,
“Fusarium spp as agents of onychomycosis in immunocompetent hosts,”
Int. J. Dermatol., 46
(8), 822
–826
(2007). http://dx.doi.org/10.1111/j.1365-4632.2007.03120.x Google Scholar
M. Halpernet al.,
“Cellulitis and nodular skin lesions due to Fusarium spp in liver transplant: case report,”
Transplant. Proc., 42
(2), 599
–600
(2010). http://dx.doi.org/10.1016/j.transproceed.2010.01.004 TRPPA8 0041-1345 Google Scholar
C. Girmeniaet al.,
“Onychomycosis as a possible origin of disseminated Fusarium solani infection in a patient with severe aplastic anemia,”
Clin. Infect. Dis., 14
(5), 1167
(1992). http://dx.doi.org/10.1093/clinids/14.5.1167 CIDIEL 1058-4838 Google Scholar
M. NucciE. Anaissie,
“Cutaneous infection by Fusarium species in healthy and immunocompromised hosts: implications for diagnosis and management,”
Clin. Infect. Dis., 35
(8), 909
–920
(2002). http://dx.doi.org/10.1086/cid.2002.35.issue-8 CIDIEL 1058-4838 Google Scholar
P. Godoyet al.,
“Onychomycosis caused by Fusarium solani and Fusarium oxysporum in São Paulo- Brazil,”
Mycopathologia, 157
(3), 287
–290
(2004). http://dx.doi.org/10.1023/B:MYCO.0000024186.32367.d4 MYCPAH 0301-486X Google Scholar
C. Romanoet al.,
“A case of primary localized cutaneous infection due to Fusarium oxysporum,”
Mycopathologia, 170
(1), 39
–46
(2010). http://dx.doi.org/10.1007/s11046-010-9290-9 MYCPAH 0301-486X Google Scholar
E. I. BoutatiE. I. Anaissie,
“Fusarium, a significant emerging pathogen in patients with hematologic malignancy: ten years’ experience at a cancer center and implications for management,”
Blood, 90
(3), 999
–1008
(1997). BLOOAW 0006-4971 Google Scholar
F. C. Oddset al.,
“Evaluation of possible correlations between antifungal susceptibilities of filamentous fungi in vitro and antifungal treatment outcomes in animal infection models,”
Antimicrob. Agents Chemother., 42
(2), 282
–288
(1998). AMACCQ 0066-4804 Google Scholar
A. Deshmukhet al.,
“Raman spectroscopy of normal oral bucal mucosa tissues: study on intact and incised biopsies,”
J. Biomed. Opt., 16
(12), 127004
(2011). http://dx.doi.org/10.1117/1.3659680 JBOPFO 1083-3668 Google Scholar
J. Kimet al.,
“In-vitro detection of artificial caries on vertical dental cavity walls using infrared photothermal radiometry and modulated luminescence,”
J. Biomed. Opt., 17
(12), 127001
(2012). http://dx.doi.org/10.1117/1.JBO.17.12.127001 JBOPFO 1083-3668 Google Scholar
S. T. Saitoet al.,
“Study of DNA-emodin interaction by FTIR and UV-vis spectroscopy,”
J. Photochem. Photobiol. B, 111
(4), 59
–63
(2012). http://dx.doi.org/10.1016/j.jphotobiol.2012.03.012 JPPBEG 1011-1344 Google Scholar
N. Tabatabaeiet al.,
“On the sensitivity of thermophotonic lock-in imaging and polarized Raman spectroscopy to early dental caries diagnosis,”
J. Biomed. Opt., 17
(2), 025002
(2012). http://dx.doi.org/10.1117/1.JBO.17.2.025002 JBOPFO 1083-3668 Google Scholar
X. Perryet al.,
“In vivo skin solvent penetration measurements using opto-thermal radiometry and fingerprint sensor,”
Int. J. Thermophys., 33
(10–11), 1787
–1794
(2012). http://dx.doi.org/10.1007/s10765-012-1318-6 IJTHDY 0195-928X Google Scholar
G. B. Junget al.,
“Effect of cross-linking with riboflavin and ultraviolet A on the chemical bonds and ultrastructure of human sclera,”
J. Biomed. Opt., 16
(12), 125004
(2011). http://dx.doi.org/10.1117/1.3662458 JBOPFO 1083-3668 Google Scholar
E. M. Moratoet al.,
“Morphological and structural changes in lung tissue infected by Paracoccidioides brasiliensis: FTIR photoacoustic spectroscopy and histological analysis,”
Photochem. Photobiol., 89
(5), 1170
–1175
(2013). http://dx.doi.org/10.1111/php.12110 PHCBAP 0031-8655 Google Scholar
A. L. M. Ubaldiniet al.,
“Fourier transform infrared photoacoustic spectroscopy study of physical chemistry interaction between human dentin and etch-&-rince adhesives in a simulated moist bond technique,”
J. Biomed. Opt., 17
(6), 065002
(2012). http://dx.doi.org/10.1117/1.JBO.17.6.065002 JBOPFO 1083-3668 Google Scholar
M. L. BaessoR. D. SnookJ. J. Andrew,
“Fourier transform infrared photoacoustic spectroscopy to study the penetration of substances through skin,”
J. Phys. IV, 4
(C7), 449
–451
(1994). http://dx.doi.org/10.1051/jp4:19947104 JPICEI 1155-4320 Google Scholar
R. D. SnookR. D. LoweM. L. Baesso,
“Photothermal spectrometry for membrane and interfacial region studies,”
Analyst, 123
(4), 587
–593
(1998). http://dx.doi.org/10.1039/a706757g ANLYAG 0365-4885 Google Scholar
G. Socrates, Biological Molecules—Macromolecules in Infrared and Raman Characteristic Group Frequencies, 330
–340 John Wiley & Son, Chichester, UK
(2001). Google Scholar
M. DiemP. R. GriffithsJ. M. Chalmers, Vibrational Spectroscopy for Medical Diagnosis, John Wiley & Son, Chichester, UK
(2008). Google Scholar
J. R. Mourantet al.,
“FTIR spectroscopy demonstrates biochemical differences in mammalian cell cultures at different growth stages,”
Biophys. J., 85
(3), 1938
–1947
(2003). http://dx.doi.org/10.1016/S0006-3495(03)74621-9 BIOJAU 0006-3495 Google Scholar
S. Xiaolianget al.,
“Detection of lung cancer tissue by attenuated total reflection-Fourier transform infrared spectroscopy—a pilot study of 60 samples,”
J. Surg. Res., 179
(1), 33
–38
(2013). http://dx.doi.org/10.1016/j.jss.2012.08.057 JSGRA2 0022-4804 Google Scholar
T. N. BhavanishankarH. P. RameshT. Shantha,
“Dermal toxicity of Fusarium toxins in combination,”
Arch. Toxicology, 61
(3), 241
–244
(1988). TXCYAC 0300-483X Google Scholar
T. Ito,
“Hair follicle is a target of stress hormone and autoimmune reactions,”
J. Dermatol. Sci., 60
(2), 67
–73
(2010). http://dx.doi.org/10.1016/j.jdermsci.2010.09.006 JDSCEI 0923-1811 Google Scholar
A. V. Marangonet al.,
“Metabolic extract of Fusarium oxysporum induces histopathologic alterations and apoptosis in the skin of Wistar rats,”
Int. J. Dermatol., 48
(7), 697
–703
(2009). http://dx.doi.org/10.1111/j.1365-4632.2009.04013.x Google Scholar
S. M. Albarenqueet al.,
“T-2 toxin-induced acute skin lesions in Wistar-derived hypotrichotic WBN/ILA-Ht rats,”
Histol. Histopathol., 14
(2), 337
–342
(1999). HIHIES 0213-3911 Google Scholar
S. M. AlbarenqueK. Doi,
“T-2 toxin-induced apoptosis in rat keratinocyte primary cultures,”
Exp. Mol. Pathol., 78
(12), 144
–149
(2005). http://dx.doi.org/10.1016/j.yexmp.2004.07.005 EXMPA6 0014-4800 Google Scholar
T. Iharaet al.,
“Apoptotic cellular damage in mice after T-2 toxin-induced acute toxicosis,”
Nat. Toxins., 5
(4), 141
–145
(1997). http://dx.doi.org/10.1002/nt.v5:4 NATOEE 1522-7189 Google Scholar
J. Shinozukaet al.,
“T-2 toxin-induced apoptosis in lymphoid organs of mice,”
Exp. Toxicol. Pathol., 49
(5), 387
–392
(1997). http://dx.doi.org/10.1016/S0940-2993(97)80124-8 ETPAEK 0940-2993 Google Scholar
S. M. Albarenqueet al.,
“Kinetics of cytokines mRNAs expression in the dorsal skin of WBN/ILA-Ht rats following topical application of T-2 toxin,”
Exp. Toxicol. Pathol., 53
(4), 271
–274
(2001). http://dx.doi.org/10.1078/0940-2993-00189 ETPAEK 0940-2993 Google Scholar
H. BigalkeA. Rummel,
“Medical aspects of toxin weapons,”
Toxicology, 214
(3), 210
–220
(2005). http://dx.doi.org/10.1016/j.tox.2005.06.015 TXCYAC 0300-483X Google Scholar
L. Marderet al.,
“Quantitative analysis of total mycotoxins in metabolic extracts of four strains of Bipolaris sorokiniana (Helminthosporium sativum),”
Proc. Biochem., 41
(1), 177
–180
(2006). http://dx.doi.org/10.1016/j.procbio.2005.06.021 PRBCAP 0032-9592 Google Scholar
R. BravoH. Macdonald-Bravo,
“Changes in nuclear distribuction of cyclin (PCNA) but not its synthesis depend on DNA replication,”
EMBO J., 4
(3), 655
–661
(1985). EMJODG 0261-4189 Google Scholar
P. Kurkiet al.,
“Expression of proliferating cell nuclear (PCNA)/cyclin during cell cycle,”
Exp. Cell Res., 166
(1), 209
–219
(1986). http://dx.doi.org/10.1016/0014-4827(86)90520-3 ECREAL 0014-4827 Google Scholar
T. HaerslevG. K. Jacobsen,
“Microwave processing for immunohistochemical demonstration of proliferating cell nuclear antigen (PCNA) in formalin-fixed and paraffin-embedded tissue,”
Acta Pathol. Microbiol. Immunol. Scand., 102
(1–6), 395
–400
(1994). http://dx.doi.org/10.1111/apm.1994.102.issue-1-6 APMSEL 0903-4641 Google Scholar
V. Kumaret al., Robbins & Cotran Pathologic Basis of Disease, 8th ed.Elsevier, Philadelphia, Pennsylvania
(2010). Google Scholar
H. K. AbbasC. J. MirochaW. T. Shier,
“Mycotoxins produced from fungi isolated from foodstuffs and soil: comparison of toxicity in fibroblasts and rat feeding tests,”
Appl. Environ. Microbiol., 48
(3), 654
–661
(1984). AEMIDF 0099-2240 Google Scholar
H. K. AbbasW. T. ShierC. J. Mirocha,
“Sensitivity of cultured human and mouse fibroblasts to trichothecenes,”
J. Assoc. Off. Anal. Chem., 67
(3), 607
–610
(1984). ANCHAM 0003-2700 Google Scholar
|