Plants respond a bit better to global warming than scientists had thought, according to a new study (AFP Photo/David McNew) More Washington (AFP) - Plants respond a bit better to global warming than scientists had thought, according to a new study that suggests their potential contribution to worsening global warming is not likely as bad as researcher believed. When it gets hotter, plants breathe harder. And the greenhouse gas carbon dioxide is produced by respiration. That's why researchers think that as Earth is warmed by CO2 from people's activities, plants may add to the emissions and make warming worse. Plants generally take in carbon dioxide during daytime photosynthesis and release carbon dioxide during respiration at night. But plants take up much more carbon dioxide in photosynthesis than they release in respiration. But now "with this new model, we predict that some ecosystems are releasing a lot less CO2 through leaf respiration than we previously thought," said coauthor Kevin Griffin, a plant physiologist at Columbia University's Lamont-Doherty Earth Observatory. The study was published Monday by the Proceedings of the National Academy of Sciences. The research found that rates of increase slow in a predictable way as temperatures rise, in every region. And the newly defined curve leads to sharply reduced estimates of respiration, especially in the coldest regions. "What we thought was a steep curve in some places is actually a little gentler," said Griffin. The biggest changes in estimates are in the coldest regions, which recently have seen warming far beyond that in temperate zones. "All of this adds up to a significant amount of carbon, so we think it's worth paying attention to," said Griffin. Lead author Mary Heskel, of Massachusetts' Marine Biological Laboratory, said the study would go far toward helping estimate "carbon storage in vegetation, and predicting concentrations of atmospheric carbon dioxide and future surface temperatures."
News Article | April 20, 2016
Sharks, skates, and rays are oddities among the fish: They have appendages growing out of the gill arch, a small cradle of bones that supports the gills. This anatomical peculiarity has led to the proposal that the paired limbs of humans, and before that the paired fins of fish, evolved from the transformation of gill arches in early fish. Genetic evidence for this theory is offered in a new study led by J. Andrew Gillis, a Royal Society University Research Fellow at the University of Cambridge, U.K., and a Whitman Center scientist at the Marine Biological Laboratory (MBL) in Woods Hole, Mass. The study, published this week in Development, demonstrates striking similarities in the genetic mechanism used to pattern gill arch appendages (called branchial rays) and fins/limbs. Studying embryos of the little skate, Gillis focused on the gene Sonic hedgehog, which produces a signaling protein whose function is well understood in the mammalian limb. Remarkably, he found that Sonic hedgehog's role in branchial rays closely parallels its role in the limb: it sets up the axis of development and, later, maintains growth of the limb skeleton. "The shared role of Sonic hedgehog in patterning branchial rays and limbs may be due to a deep evolutionary relationship between the two," Gillis says, "or it may simply be that two unrelated appendages independently use the same gene for the same function." Ongoing studies comparing the function of other genes during branchial ray and fin/limb development will help to resolve this. Gillis will continue his research at the MBL this summer using skates collected and supplied by the Marine Resources Department. "Branchial rays will figure prominently in the story of the evolutionary origin of vertebrate animal appendages, either by shedding light on the evolutionary antecedent of paired fins/limbs, or by teaching us about the genetic mechanisms that animals can use to invent new appendages," Gillis says.
All non-transgenic, transgenic and control mice used in this study were derived from in house breeding colonies backcrossed > 12 generations onto C57/BL6 backgrounds. All mice used were young adult females between two and four months old at the time of spinal cord injury. All transgenic mice used have been previously well characterized or are the progeny of crossing well-characterized lines: (1) mGFAP-TK transgenic mice line 7.115, 16, 49; (2) mGFAP-Cre-STAT3-loxP mice generated by crossing STAT3-loxP mice with loxP sites flanking exon 22 of the STAT3 gene50 with mGFAP-Cre mice line 73.1217, 18; (3) loxP-STOP-loxP-DTR (diphtheria toxin receptor) mice21; (4) mGFAP-Cre-RiboTag mice generated by crossing mice with loxP-STOP-loxP-Rpl22-HA (RiboTag)26 with mGFAP-Cre mice line 73.1217, 18; (5) loxP-STOP-loxP-tdTomato reporter mice51. All mice were housed in a 12-h light/dark cycle in a specific-pathogen-free facility with controlled temperature and humidity and were allowed free access to food and water. All experiments were conducted according to protocols approved by the Animal Research Committee of the Office for Protection of Research Subjects at University of California, Los Angeles. All surgeries were performed under general anaesthesia with isoflurane in oxygen-enriched air using an operating microscope (Zeiss, Oberkochen, Germany), and rodent stereotaxic apparatus (David Kopf, Tujunga, CA). Laminectomy of a single vertebra was performed and severe crush spinal cord injuries (SCI) were made at the level of T10 using No. 5 Dumont forceps (Fine Science Tools, Foster City, CA) without spacers and with a tip width of 0.5 mm to completely compress the entire spinal cord laterally from both sides for 5 s16, 17, 18. For pre-conditioning lesions, sciatic nerves were transected and ligated one week before SCI. Hydrogels were injected stereotaxically into the centre of SCI lesions 0.6 mm below the surface at 0.2 μl per minute using glass micropipettes (ground to 50–100 μm tips) connected via high-pressure tubing (Kopf) to 10-μl syringes under control of microinfusion pumps, two days after SCI52. Tract tracing was performed by injection of biotinylated dextran amine 10,000 (BDA, Invitrogen) 10% wt/vol in sterile saline injected 4 × 0.4 μl into the left motor cerebral cortex 14 days before perfusion to visualize corticospinal tract (CST) axons, or choleratoxin B (CTB) (List Biological Laboratory, Campbell, CA) 1 μl of 1% wt/vol in sterile water injected into both sciatic nerves three days before perfusion to visualize ascending sensory tract (AST) axons33. AAV2/5-GfaABC1D-Cre (see below) was injected either 3 or 6 × 0.4 μl (1.29 × 1013 gc ml−1 in sterile saline) into and on either side of mature SCI lesions two weeks after SCI, or into uninjured spinal cord after T10 laminectomy. All animals received analgesic before wound closure and every 12 h for at least 48 h post-injury. Animals were randomly assigned numbers and evaluated thereafter blind to genotype and experimental condition. Adeno-associated virus 2/5 (AAV) vector with a minimal GFAP promoter (AAV2/5 GfaABC1D) was used to target Cre-recombinase expression selectively to astrocytes53, 54, 55. Diblock co-polypeptide hydrogel (DCH) K L was fabricated, tagged with blue fluorescent dye (AMCA-X) and loaded with growth factor and antibody cargoes as described38, 39, 52. Cargo molecules comprised: human recombinant NT3 and BDNF were gifts (Amgen, Thousand Oaks, CA, (NT3 Lot#2200F4; BDNF Lot#2142F5A) or were purchased from PeproTech (Rocky Hill, NJ; NT3 405-03, Lot#060762; BDNF 405-02 Lot#071161). Function blocking anti-CD29 mouse monoclonal antibody was purchased from BD Bioscience (San Diego, CA) as a custom order at 10.25 mg ml−1 (product #BP555003; lot#S03146). Freeze dried K L powder was reconstituted on to 3.0% or 3.5% wt/vol basis in sterile PBS without cargo or with combinations of NT3 (1.0 μg μl−1), BDNF (0.85 μg μl−1) and anti-CD29 (5 μg μl−1). DCH mixtures were prepared to have G′ (storage modulus at 1 Hz) between 75 and 100 Pascal (Pa), somewhat below that of mouse brain at 200 Pa (refs 38, 39). GCV (Cytovene-IV Hoffman LaRoche, Nutley, NJ), 25 mg kg−1 per day dissolved in sterile physiological saline was administered as single daily subcutaneous injections starting immediately after surgery and continued for the first 7 days after SCI. Bromodeoxyuridine (BrdU, Sigma), 100 mg kg−1 per day dissolved in saline plus 0.007 M NaOH, was administered as single daily intraperitoneal injections on days 2 through 7 after SCI. Diphtheria toxin A (DT, Sigma #DO564) 100 ng in 100 μl sterile saline was administered twice daily as intraperitoneal injections for ten days starting three weeks after injection of AAV2/5-GfaABC1D-Cre to loxP-DTR mice (which was 5 weeks after SCI) (see timeline in Extended Data Fig. 1d). Two days after SCI, all mice were evaluated in open field and mice exhibiting any hindlimb movements were not studied further. Mice that passed this pre-determined inclusion criterion were randomized into experimental groups for further treatments and were thereafter evaluated blind to their experimental condition. At 3, 7, 14 days and then weekly after SCI, hindlimb movements were scored using a simple six-point scale in which 0 is no movement and 5 is normal walking17. After terminal anaesthesia by barbiturate overdose mice were perfused transcardially with 10% formalin (Sigma). Spinal cords were removed, post-fixed overnight, and cryoprotected in buffered 30% sucrose for 48 h. Frozen sections (30 μm horizontal) were prepared using a cryostat microtome (Leica) and processed for immunofluorescence as described16, 17, 18. Primary antibodies were: rabbit anti-GFAP (1:1,000; Dako, Carpinteria, CA); rat anti-GFAP (1:1,000, Zymed Laboratories); goat anti-CTB (1:1,000, List Biology Lab); rabbit anti-5HT (1:2,000, Immunostar); goat anti-5HT (1:1,000, Immunostar); mouse anti-CSPG22 (1:100, Sigma); rabbit-anti haemagglutinin (HA) (1:500 Sigma); mouse-anti HA (1:3,000 Covance); sheep anti-BrdU (1:6,000, Maine Biotechnology Services, Portland, ME); rabbit anti-laminin (1:80, Sigma, Saint Louis, MO); guinea pig anti-NG2 (CSPG4) (E. G. Hughes and D. W. Bergles56, Baltimore, MA); goat anti-aggrecan (1:200, NOVUS); rabbit anti-brevican (1:300, NOVUS); mouse anti-neurocan (1:300, Milipore); mouse anti-phosphacan (1:500, Sigma); goat anti-versican (1:200, NOVUS); rabbit anti-neurglycan C (CSPG5) (1:200, NOVUS). Fluorescence secondary antibodies were conjugated to: Alexa 488 (green) or Alexa 350 (blue) (Molecular Probes), or to Cy3 (550, red) or Cy5 (649, far red) all from (Jackson Immunoresearch Laboratories). Mouse primary antibodies were visualized using the Mouse-on-Mouse detection kit (M.O.M., Vector). BDA tract-tracing was visualized with streptavidin-HRP plus TSB Fluorescein green or Tyr-Cy3 (Jackson Immunoresearch Laboratories). Nuclear stain: 4′,6′-diamidino-2-phenylindole dihydrochloride (DAPI; 2 ng ml−1; Molecular Probes). Sections were coverslipped using ProLong Gold anti-fade reagent (InVitrogen, Grand Island, NY). Sections were examined and photographed using deconvolution fluorescence microscopy and scanning confocal laser microscopy (Zeiss, Oberkochen, Germany). Axons labelled by tract tracing or immunohistochemistry were quantified using image analysis software (NeuroLucida, MicroBrightField, Williston, VT) operating a computer-driven microscope regulated in the x, y and z axes (Zeiss) by observers blind to experimental conditions. Using NeuroLucida, lines were drawn across horizontal spinal cord sections at SCI lesion centres and at regular distances on either side (Fig. 1a) and the number of axons intercepting lines was counted at 63× magnification under oil immersion by observers blind to experimental conditions. Similar lines were drawn and axons counted in intact axon tracts 3 mm proximal to SCI lesions and the numbers of axon intercepts in or near lesions were expressed as percentages of axons in the intact tracts in order to control for potential variations in tract-tracing efficacy or intensity of immunohistochemistry among animals. Two sections at the level of the CST or AST, and three sections through the middle of the cord for 5HT, were counted per mouse and expressed as total intercepts per location per mouse. To determine efficacy of axon transection after SCI, we examined labelling 3 mm distal to SCI lesion centres, with the intention of eliminating mice that had labelled axons at this location on grounds that these mice may have had incomplete lesions. However, all mice that had met the strict behavioural inclusion criterion of no hindlimb movements two days after severe crush SCI, exhibited no detectable axons 3 mm distal to SCI lesions regardless of treatment group. Sections stained for GFAP, CSPG or laminin were photographed using constant exposure settings. Single-channel immunofluorescence images were converted to black and white and thresholded (Fig. 1d and Extended Data Fig. 2b) and the amount of stained area measured in different tissue compartments using NIH ImageJ software. Areas are shown in graphs as mean values plus or minus standard error of the means (s.e.m.). Statistical evaluations of repeated measures were conducted by ANOVA with post hoc, independent pairwise analysis as per Newman-Keuls (Prism, GraphPad, San Diego, CA). Power calculations were performed using G*Power Software v188.8.131.52 (ref. 57). For quantification of histologically derived neuroanatomical outcomes such as numbers of axons or percentage of area stained for GFAP or CSPG, group sizes were used that were calculated to provide at least 80% power when using the following parameters: probability of type I error (α) = 0.05, a conservative effect size of 0.25, 2–8 treatment groups with multiple measurements obtained per replicate. Using Fig. 5j as an example, evaluation of n = 5 biological replicates (with multiple measurements per replicate) in each of 8 treatment groups provided greater than 88% power. For dot blot immunoassay of chondroitin sulfate proteoglycans (CSPG), spinal cord tissue blocks were lysed and homogenized in standard RIPA (radio-immunoprecipitation assay) buffer. LDS (lithium dodecyl sulfate) buffer (Life Technologies) was added to the post-mitochondrial supernatant and 2 μl containing 2 μg μl−1 protein was spotted onto a nitrocellulose membrane (Life Technologies), set to dry and incubated overnight with mouse anti-chondroitin sulfate antibody (CS56, 1:1000, Sigma Aldrich), an IgM-monoclonal antibody that detects glyco-moieties of all CSPGs22. CS56 immunoreactivity was detected on X-ray film with alkaline phosphatase-conjugated secondary antibody and chemiluminescent substrate (Life Technologies). Densitometry measurements of CS56 immunoreactivity were obtained using ImageJ software (NIH) and normalized to total protein (Poncau S) density58. Densities are shown in graphs as mean values plus or minus standard error of the means (s.e.m.). Two weeks after SCI, spinal cords of wild-type control (GFAP-RiboTag) and STAT3-CKO (GFAP-STAT3CKO-RiboTag) mice were rapidly dissected out of the spinal canal. The central 3 mm of the lower thoracic lesion including the lesion core and 1 mm rostral and caudal were then rapidly removed and snap frozen in liquid nitrogen. Haemagglutinin (HA) immunoprecipitation (HA-IP) of astrocyte ribosomes and ribosome-associated mRNA (ramRNA) was carried out as described26. The non-precipitated flow-through (FT) from each IP sample was collected for analysis of non-astrocyte total RNA. HA and FT samples underwent on-column DNA digestion using the RNase-Free Dnase Set (Qiagen) and RNA purified with the RNeasy Micro kit (Qiagen). Integrity of the eluted RNA was analysed by a 2100 Bioanalyzer (Agilent) using the RNA Pico chip, mean sample RIN = 8.0 ± 0.95. RNA concentration determined by RiboGreen RNA Assay kit (Life Technologies). cDNA was generated from 5 ng of IP or FT RNA using the Nugen Ovation 2 RNA-Seq Sytstem V2 kit (Nugen). 1 μg of cDNA was fragmented using the Covaris M220. Paired-end libraries for multiplex sequencing were generated from 300 ng of fragmented cDNA using the Apollo 324 automated library preparation system (Wafergen Biosystems) and purified with Agencourt AMPure XP beads (Beckman Coulter). All samples were analysed by an Illumina NextSeq 500 Sequencer (Illumina) using 75-bp paired-end sequencing. Reads were quality controlled using in-house scripts including picard-tools, mapped to the reference mm10 genome using STAR59, and counted using HT-seq60 with mm10 refSeq as reference, and genes were called differentially expressed using edgeR61. Individual gene expression levels in the Fig. 4e histogram are shown as mean FPKM (fragments per kilobase of transcript sequence per million mapped fragments). Additional details of differential expression analysis are described in the legends of Fig. 4 and Extended Data Figs 3 and 4. Raw and normalized data have been deposited in the NCBI Gene Expression Omnibus and are accessible through accession number GSE76097. To ensure the widespread distribution of these datasets, we have created a user-friendly website that enables searching for individual genes of interest https://astrocyte.rnaseq.sofroniewlab.neurobio.ucla.edu.
News Article | April 7, 2016
In just about every nook, cranny, and crevice of our planet, some sort of life manages to thrive—whether it’s under an Antarctic ice sheet, in super-salty Arctic water, or in Chile’s Atacama desert, one of the driest and harshest environments in the world. A US scientist has found something living in another surprising place: in the rocky sediment deep under the Atlantic Ocean, 50 to 250 meters beneath the seafloor, which is itself under 4.5 km—that’s more than 2.7 miles—of ocean water. With no sunlight and few nutrients, not to mention extreme pressure, you won’t find fish or many other creatures that deep. These tiny microbes can eke out a living in deep ocean sediment and rock. Learning about them could help us find life in bizarre environments on other planets, too. In the new paper, Julie Huber, associate scientist at the Marine Biological Laboratory in Woods Hole, Mass., describes the microbial community she and her team found way out at the bottom of the Atlantic. “It’s the middle of the ocean,” she told Motherboard. “Water, as far as the eye can see.” At North Pond in the mid-Atlantic, the ocean crust is young, and its circulating fluids are cold. Photo: Marine Biological Laboratory Deep down underneath all that water is a sediment pocket called North Pond, on the western edge of the Mid-Atlantic Ridge. That’s where new ocean crust is being formed as plates push apart. That crust isn’t static, she continued. “Fluids are still moving through it,” as seawater rushes through its crevices. Samples were collected with the Integrated Ocean Drilling Program, an international project that drills deep into the seafloor for science—not the same as offshore oil drilling, although technologies have been swapped between the two. For all the strangeness of the environment they live in, these microbes aren’t necessarily “extremophiles,” Huber said. “They appear to be closely related to [others found] in seawater. But we’re finding genetic signatures suggesting they are slightly different,” in ways that aren’t yet understood. She and others are trying to piece that together now, studying their DNA. Huber wasn’t surprised to learn that something could live in a “cold crustal aquifer,” as she calls this deep-ocean environment. (Most of her other work focuses on high-temperature hydrothermal vents and underwater volcanoes.) “It’s pretty rare not to find microbes,” she said. “They seem to find a way.” Given that life takes hold basically everywhere on our planet, could we find it on another one? Scientists are excited by the idea that they could one day find something living on Enceladus, an icy moon of Saturn. “Based on modelling, it looks like pretty much the only energy available there is methane and carbon dioxide,” maybe a bit of hydrogen, said Huber, who’s received NASA funding for some of her research. “Only a handful of microbes can use those on our planet, and they’re pretty specialized.” By studying life in lower-energy environments—like the rock and sediment at the bottom of the ocean—we’ll learn more about what tricks microbes use to eke out a living. Hopefully, it’s preparation for one day getting to Enceladus.
Protected areas such as rainforests occupy more than one-tenth of the Earth’s landscape, and provide invaluable ecosystem services, from erosion control to pollination to biodiversity preservation. They also draw heat-trapping carbon dioxide (CO ) from the atmosphere and store it in plants and soil through photosynthesis, yielding a net cooling effect on the planet. Determining the role protected areas play as carbon sinks — now and in decades to come — is a topic of intense interest to the climate-policy community as it seeks science-based strategies to mitigate climate change. Toward that end, a study in the journal Ambio estimates for the first time the amount of CO sequestered by protected areas, both at present and throughout the 21st century as projected under various climate and land-use scenarios. Based on their models and assuming a business-as-usual climate scenario, the researchers projected that the annual carbon sequestration rate in protected areas will decline by about 40 percent between now and 2100. Moreover, if about one-third of protected land is converted to other uses by that time, due to population and economic pressures, carbon sequestration in the remaining protected areas will become negligible. “Our study highlights the importance of protected areas in slowing the rate of climate change by pulling carbon dioxide out of the atmosphere and sequestering it in plants and soils, especially in forested areas,” said Jerry Melillo, the study’s lead author. Melillo is a distinguished scientist at the Marine Biological Laboratory (MBL) in Woods Hole, Massachusetts, and former director of the MBL’s Ecosystems Center. “Maintaining existing protected areas, enlarging them and adding new ones over this century are important ways we can manage the global landscape to help mitigate climate change.” Based on a global database of protected areas, a reconstruction of global land-use history, and a global biogeochemistry model, the researchers estimated that protected areas currently sequester 0.5 petagrams (500 billion kilograms) of carbon each year, or about 20 percent of the carbon sequestered by all land ecosystems annually. Using an integrated modeling framework developed by the MIT Joint Program on the Science and Policy of Global Change, they projected that under a rapid climate-change scenario that extends existing climate policies; keeps protected areas off-limits to development; and assumes continued economic growth and a 1 percent annual increase in agricultural productivity, the annual carbon sequestration rate in protected areas would fall to about 0.3 petagrams of carbon by 2100. When they ran the same scenario but allowed for possible development of protected areas, they projected that more than one-third of today’s protected areas would be converted to other uses. This would reduce carbon sequestration in the remaining protected areas to near zero by the end of the century. (The protected areas that are not converted would be the more marginal systems that have low productivity, and thus low capacity to sequester carbon.) Based on this analysis, the researchers concluded that unless current protected areas are preserved and expanded, their capacity to sequester carbon will decline. The need for expansion is driven by climate change: As the average global temperature rises, so, too, will plant and soil respiration in protected and unprotected areas alike, thereby reducing their ability to store carbon and cool the planet. “This work shows the need for sufficient resources dedicated to actually prevent encroachment of human activity into protected areas,” said John Reilly, one of the study’s coauthors and the co-director of the MIT Joint Program on the Science and Policy of Global Change. The study was supported by the David and Lucille Packard foundation, the National Science Foundation, the U.S. Environmental Protection Agency, and the U.S. Department of Energy.