Chemometric Analysis Applied In 1h Hr-mas Nmr And Ft-ir Data For Chemotaxonomic Distinction Of Intact Lichen Samples

This paper describes the potentiality of chemometric analysis applied in 1H HR-MAS NMR and FT-IR data for lichen chemotaxonomic investigations. Lichens present a difficult morphologic differentiation and the chemical analyses are frequently employed for their taxonomic classification, mainly due to the secondary metabolites to be relatively constant for these organisms. The lichen chemotaxonomic classification is usually carried out by color reactions, chromatography, fluorescence and mass spectrometry analysis, where the identification is obtained by one or more techniques. There are some papers which use the carbohydrate content in chemotaxonomy investigation. However, the majority of these techniques involve laborious and time consuming sample pre-treatment. This work focuses on application of 1H high resolution magic angle spinning - nuclear magnetic resonance (HR-MAS NMR) and Fourier transform infrared (FT-IR) associated with chemometric analysis to intact samples. In comparison to other traditional techniques, 1H HR-MAS NMR and FT-IR allied with chemometrics provided a fast and economic method for lichen chemotaxonomy. Both methods were useful for lichen analysis and permitted the satisfactory distinction among families, genera and species, although better results were achieved for FT-IR data. © 2007 Elsevier B.V. All rights reserved.5951-2 SPEC. ISS.38Nash III, T.H., (1996) Lichen biology, , Cambridge University Press, Cambridge 303 pQuilhot, W., Leighton, G., Flores, E., Fernandes, E., Pena, W., Guzman, G., (1987) Acta Farm. Bonaerense, 6 (1), pp. 15-22Eifler-Lima, V.L., Sperry, A., Sinbandhit, S., Boustie, J., Tomasi, S., Schenkel, E., (2000) Magn. Reson. Chem., 38, pp. 472-474Honda, N.K., Vilegas, W., (1998) Quím. Nova, 21 (6), pp. 110-125Carbonero, E.R., Sassaki, G.L., Stuelp, P.M., Gorin, P.A.J., Woranovicz-Barreira, S.M., Iacomini, M., (2001) FEMS Microbiol. Lett., 194 (1), pp. 65-69Yokota, I., Shibata, S., Saitô, S., (1979) Carbohydr. Res., 69, pp. 252-258Teixeira, A.Z.A., Iacomini, M., Gorin, P.A., (1995) Carbohydr. Res., 266 (2), pp. 309-314Stuelp, P.M., Carneiro Leão, A.M.A., Gorin, P.A.J., Iacomini, M., (1999) Carbohydr. Pol., 40, pp. 101-106Carbonero, E.R., Montai, A.V., Stuelp, P.M., Woranovicz-Barreira, S.M., Gorin, P.A.J., Iacomini, M., (2002) Phytochem., 61, pp. 681-686Carbonero, E.R., Montai, A.V., Mellinger, C.G., Eliasaro, S., Sassaki, G.L., Gorin, P.A.J., Iacomini, M., (2005) Phytochem., 66, pp. 929-934Hale Jr., M.E., (1983) The biology of lichens. third ed., , Edward Arnold, Baltimore 190 pCheng, L.L., Chang, I., Smith, B.L., Gonzalez, R.G., (1998) J. Magn. Res., 135, pp. 194-202Moka, D., Vorreuther, R., Schicha, H., Spraul, M., Humpfer, E., Lipinski, M., Foxall, P.J.D., Lindon, J.C., (1997) Anal. Commun., 34, pp. 107-109Gil, A.M., Duarte, I.F., Delgadillo, I., Colquhoun, I.J., Casuscelli, F., Humpfer, E., Spraul, M., (2000) J. Agric. Food Chem., 48, pp. 1524-1536Ni, Q.X., Eads, T.M., (1993) J. Agric. Food Chem., 41, pp. 1035-1040Broberg, A., Kenne, L., Pedersen, M., (1998) Planta, 206 (2), pp. 300-307Brescia, M.A., Di Martino, G., Fares, C., Di Fonzo, N., Platani, C., Chelli, S., Reniero, F., Sacco, A., (2002) Cereal Chem., 79 (2), pp. 238-242Sacco, A., Bolsi, I.N., Massini, R., Spraul, M., Humpfer, E., Cheli, S., (1998) J. Agric. Food Chem., 46, pp. 4242-4249Luginbühl, W., Jimeno, J., Zehntner, U., (2006) LWT, 39, pp. 152-158Reid, L.M., O'Donnell, C.P., Downey, G., Trends Food Sci. (2006) Tech., 17, pp. 344-353Schneider, R., Charrier, F., Moutounet, M., Baumes, R., (2004) Anal. Chim. Acta, 513, pp. 91-96Fischer, G., Braun, S., Thissen, R., Dott, W., (2006) J. Microbiol. Methods, 64, pp. 63-77Belton, P.S., Colquhoum, I.J., Kemsley, E.K., Delgadillo, I., Roma, P., Dennis, M.J., Sharman, M., Spraul, M., (1998) Food Chem., 61 (2), pp. 207-213Lai, Y.W., Kemsley, E.K., Wilson, R.H., (1994) J. Agric. Food Chem., 42, pp. 1154-1159Defernez, M., Kemsley, E.K., Wilson, R.H., (1996) J. Agric. Food Chem., 43, pp. 109-113Vogels, J.T.W.E., Terwel, L., Tas, A.C., Van den Berg, F., Dukel, F., Van der Greef, J., (1996) J. Agric. Food Chem., 44, pp. 175-180Ward, J.L., Harris, C., Lewis, J., Beale, M.H., (2003) Phytochem., 62, pp. 949-957Howells, S.L., Maxwell, R.J., Peet, A.C., Griffiths, J.R., (1992) Magn. Reson. Med., 28, pp. 214-236Holmes, E., Tsang, T.M., Tabrizi, S.J., (2006) J. Am. Soc. Exp. NeuroTher., 3, pp. 358-372Szabo de Edelenyi, F., Simonetti, A.W., Postma, G., Huo, R., Buydens, L.M.C., (2005) Anal. Chim. Acta, 544, pp. 36-46Kowalski, B.R., Bender, C.F., (1972) Anal. Chem., 44 (8), pp. 1405-141

Fenologia de Lafoensia pacari A.St.-Hil. (Lythraceae) em Barra do Garças, Mato Grosso, Brasil Phenology of Lafoensia pacari A.St.-Hil. (Lythraceae) in Barra do Garças, Mato Grosso State, Brazil

Lafoensia pacari A.St.-Hil. é uma espécie da flora do cerrado usada na medicina popular como anti-úlcera, antifúngica, antibactericida, anti-inflamatória, febrífuga, para emagrecimento e no tratamento de pneumonia, dores de estômago e coceiras. Estudou-se a fenologia de L. pacari no cerrado do Parque Estadual da "Serra Azul", região de Barra do Garças-MT (15º 51' 58" S e 52º 15' 37" W), à 645 m de altitude, durante o período de 24 meses. Verificou-se que as fenofases são sazonais, com floração nos meses de abril a agosto, frutificação de junho a setembro, brotação no início da estação chuvosa de outubro a dezembro e queda de folhas de julho a setembro, no final da estação seca.<br>Lafoensia pacari is a species from the Brazilian cerrado used in folk medicine to control ulcers, fungal and bacterial diseases, inflammations, fevers, pneumonia, stomachaches, and itching, as well as to lose weight. L. pacari phenology was studied for 24 months in the cerrado at "Serra Azul" State Park, in the region of Barra do Garças, Mato Grosso State, Brazil (15º 51' 58" S and 52º 15' 37" W), at 645 m altitude. Phenophases are seasonal, with flowering from April to August, fruiting from June to September, sprouting from October to December, during the beginning of the rainy season, and leaf fall from July to September, at the end of the dry season

Atmosphere Impact Losses

Determining the origin of volatiles on terrestrial planets and quantifying atmospheric loss during planet formation is crucial for understanding the history and evolution of planetary atmospheres. Using geochemical observations of noble gases and major volatiles we determine what the present day inventory of volatiles tells us about the sources, the accretion process and the early differentiation of the Earth. We further quantify the key volatile loss mechanisms and the atmospheric loss history during Earth’s formation. Volatiles were accreted throughout the Earth’s formation, but Earth’s early accretion history was volatile poor. Although nebular Ne and possible H in the deep mantle might be a fingerprint of this early accretion, most of the mantle does not remember this signature implying that volatile loss occurred during accretion. Present day geochemistry of volatiles shows no evidence of hydrodynamic escape as the isotopic compositions of most volatiles are chondritic. This suggests that atmospheric loss generated by impacts played a major role during Earth’s formation. While many of the volatiles have chondritic isotopic ratios, their relative abundances are certainly not chondritic again suggesting volatile loss tied to impacts. Geochemical evidence of atmospheric loss comes from the He3/22Ne, halogen ratios (e.g., F/Cl) and low H/N ratios. In addition, the geochemical ratios indicate that most of the water could have been delivered prior to the Moon forming impact and that the Moon forming impact did not drive off the ocean. Given the importance of impacts in determining the volatile budget of the Earth we examine the contributions to atmospheric loss from both small and large impacts. We find that atmospheric mass loss due to impacts can be characterized into three different regimes: 1) Giant Impacts, that create a strong shock transversing the whole planet and that can lead to atmospheric loss globally. 2) Large enough impactors (mcap≳2ρ0(πhR)3/2, rcap∼25km for the current Earth), that are able to eject all the atmosphere above the tangent plane of the impact site, where h, R and ρ0 are the atmospheric scale height, radius of the target, and its atmospheric density at the ground. 3) Small impactors (mmin&gt; 4 πρ0h3, rmin∼1km for the current Earth), that are only able to eject a fraction of the atmospheric mass above the tangent plane. We demonstrate that per unit impactor mass, small impactors with rmin&lt; r&lt; rcap are the most efficient impactors in eroding the atmosphere. In fact for the current atmospheric mass of the Earth, they are more than five orders of magnitude more efficient (per unit impactor mass) than giant impacts, implying that atmospheric mass loss must have been common. The enormous atmospheric mass loss efficiency of small impactors is due to the fact that most of their impact energy and momentum is directly available for local mass loss, where as in the giant impact regime a lot of energy and momentum is ’wasted’ by having to create a strong shock that can transverse the entirety of the planet such that global atmospheric loss can be achieved. In the absence of any volatile delivery and outgassing, we show that the population of late impactors inferred from the lunar cratering record containing 0.1%&nbsp;M⊕ is able to erode the entire current Earth’s atmosphere implying that an interplay of erosion, outgassing and volatile delivery is likely responsible for determining the atmospheric mass and composition of the early Earth. Combining geochemical observations with impact models suggest an interesting synergy between small and big impacts, where giant impacts create large magma oceans and small and larger impacts drive the atmospheric loss