Biodegradation of synthetic polymers in soils: Tracking carbon into CO2 and microbial biomass

http://advances.sciencemag.org/content/4/7/eaas9024
Biodegradation of synthetic polymers in soils: Tracking carbon into CO2 and microbial biomass

1. Michael Thomas Zumstein¹,*,Arno Schintlmeister²,³,Taylor Frederick Nelson¹,Rebekka Baumgartner¹, Dagmar Woebken²,Michael Wagner²,³,Hans-Peter E. Kohler⁴,Kristopher McNeill¹ and Michael Sander¹,†

Abstract

Plastic materials are widely used in agricultural applications to achieve food security for the growing world population. The use of biodegradable instead of nonbiodegradable polymers in single-use agricultural applications, including plastic mulching, promises to reduce plastic accumulation in the environment. We present a novel approach that allows tracking of carbon from biodegradable polymers into CO₂ and microbial biomass. The approach is based on ¹³C-labeled polymers and on isotope-specific analytical methods, including nanoscale secondary ion mass spectrometry (NanoSIMS). Our results unequivocally demonstrate the biodegradability of poly(butylene adipate-co-terephthalate) (PBAT), an important polyester used in agriculture, in soil. Carbon from each monomer unit of PBAT was used by soil microorganisms, including filamentous fungi, to gain energy and to form biomass. This work advances both our conceptual understanding of polymer biodegradation and the methodological capabilities to assess this process in natural and engineered environments.

INTRODUCTION

Modern agriculture heavily relies on the use of plastic materials in various applications, a practice coined plasticulture. Mulching with plastic films is a major application with a global market volume of approximately 2 × 10⁶ tons per year (1). Mulch films are placed onto agricultural soils to improve conditions for plant growth while lowering consumption of water, herbicides, and fertilizer and also minimizing soil erosion (1, 2). Because of these benefits, mulching with plastic films helps to secure food for the growing world population. However, mulch films are commonly composed of nonbiodegradable polyethylene and, therefore, accumulate in agricultural soils and surrounding receiving environments if incompletely retrieved after use. These accumulations have negative ecologic and economic impacts, including decreased soil productivity (3–5). A promising strategy to overcome these risks is to use mulch films composed of polymers that biodegrade in soils (1, 6–8).

Biodegradation of polymers requires microorganisms to metabolize all organic components of the polymer. Biodegradation in soil involves several key steps (Fig. 1): (i) colonization of the polymer surface by microorganisms, (ii) secretion of extracellular microbial enzymes that depolymerize the polymer into low–molecular weight compounds, and (iii) microbial uptake and utilization of these compounds, incorporating polymer carbon into biomass or releasing it as CO₂ (9).

Fig. 1 Key steps in the biodegradation of polymers in soils.

Microorganisms colonize the polymer surface and secrete extracellular enzymes that depolymerize the polymer. The formed low–molecular weight hydrolysis products are taken up by the microorganisms and used both for energy production, resulting in the formation of CO₂, and for the synthesis of cellular structures and macromolecules, resulting in incorporation of polymer-derived carbon into the microbial biomass. The boxes on the right depict the analytical methods we used to study these steps. NMR, nuclear magnetic resonance.

Fig. 1 Key steps in the biodegradation of polymers in soils.Microorganisms colonize the polymer surface and secrete extracellular enzymes that depolymerize the polymer. The formed low–molecular weight hydrolysis products are taken up by the microorganisms and used both for energy production, resulting in the formation of CO₂, and for the synthesis of cellular structures and macromolecules, resulting in incorporation of polymer-derived carbon into the microbial biomass. The boxes on the right depict the analytical methods we used to study these steps. NMR, nuclear magnetic resonance.Here, we examined each of the above steps for poly(butylene adipate-co-terephthalate) (PBAT), an aliphatic-aromatic statistical copolyester of large importance in the market of biodegradable mulch films (7). While previous studies provided indirect indications for PBAT biodegradation in soils based on determining PBAT mass loss and changes in its physicochemical properties (10–12), we here use a novel workflow using stable carbon isotope-labeled PBAT to directly and unequivocally demonstrate its biodegradation in soil (table S1). This workflow included incubation of ¹³C-labeled polymer films in soil with continuous quantification of polymer-derived ¹³CO₂ by isotope-specific cavity ring-down spectroscopy (CRDS) (13). The use of ¹³C-labeled polymers allowed us to distinguish polymer-derived CO₂ from CO₂formed by soil organic matter mineralization. After incubation, we imaged the polymer film surfaces using scanning electron microscopy (SEM) and demonstrated the incorporation of polymer-derived ¹³C into the biomass of film-colonizing microorganisms using element-specific, isotope-selective nanoscale secondary ion mass spectrometry (NanoSIMS) (14). We studied three PBAT variants that had similar physicochemical properties and comparable total ¹³C contents, but varied in the monomer that contained the ¹³C-label [that is, butanediol (P*BAT), adipate (PB*AT), or terephthalate (PBA*T)] (Fig. 2A and table S2). The use of these variants allowed us to follow biodegradation of all PBAT building blocks. The presented workflow is a novel approach to study the fundamental steps in polymer biodegradation in complex systems (15–17).

DISCUSSION

This work presents an experimental approach to study polymer biodegradation in soils and to assess the key steps involved in this process: microbial polymer colonization, enzymatic depolymerization on the polymer surface, and microbial uptake and utilization of the released low–molecular weight compounds. Central to the approach is the use of polymer variants that are ¹³C-labeled in all monomer units of the polymer, thereby allowing us to assess whether all organic components of the polymer material are used by soil microorganisms. The label further allows tracing of polymer-derived carbon into both CO₂ and microbial biomass. Using this approach, we demonstrate here the biodegradability of PBAT in soil. Biodegradability renders PBAT a more environmentally friendly alternative to persistent polymer materials for use in plasticulture, including single-use applications such as plastic mulching. Our results further imply that incorporation of polymer-derived carbon into microbial biomass needs to be taken into consideration in regulatory guidelines for determining biodegradability of polymers. Currently, these guidelines are solely based on extents of CO₂ formation. Furthermore, the finding of subcellular structures within PBAT-colonizing fungi highly enriched in polymer-derived carbon might represent compartments in which carbon is stored (for example, in the form of neutral lipids) when fungi are limited by the availability of nutrients other than carbon (22). These limitations are plausible for microorganisms growing on PBAT and other polymers that do not contain nitrogen and phosphorous. If these limitations occur, increasing the availability of soil nutrients to microorganisms colonizing the polymer surface is expected to enhance polymer biodegradation.

This work demonstrates PBAT biodegradation in a selected agricultural soil over 6 weeks of incubation. Future studies extending on this work will need to assess variations in the rates and extents of PBAT mineralization among different agricultural soils, also over longer-time incubations. Furthermore, we propose studies that are directed toward identifying soil microorganisms that are actively involved in PBAT biodegradation. While the NanoSIMS-based approach presented here allows us to unambiguously demonstrate incorporation of polyester carbon into soil microbial biomass, it is not a high-throughput technique. Alternative approaches, including the extraction of targeted biomolecules from soils containing ¹³C-labeled polymers followed by quantifying the ¹³C contents in the extracted molecules, will allow us to analyze larger sample sets and thereby to systematically determine potential variations among soil microorganisms in the extent to which they incorporate polymer-derived carbon into their biomass.

MATERIALS AND METHODS

Experimental design

The objective of this study was to develop an experimental approach to demonstrate biodegradation of PBAT in an agricultural soil. As biodegradation includes mineralization of PBAT carbon to CO₂, as well as the incorporation of PBAT-derived carbon into the biomass of soil microorganisms, we addressed both of these processes in controlled laboratory experiments. We followed PBAT mineralization during soil incubation using an isotope-specific CRDS for the quantification of formed CO₂. For each of the three PBAT variants, we simultaneously incubated seven films in one incubation bottle filled with soil to allow precise quantification of PBAT mineralization to CO₂. The soil incubations were terminated after 6 weeks (that is, when approximately 10% of the PBAT carbon had been mineralized) to ensure that PBAT films were still intact for the subsequent imaging analyses. We revealed incorporation of PBAT-derived carbon into biomass using NanoSIMS, which enabled identification of subcellular features and determination of the carbon isotope composition of the PBAT film surface and the colonizing microorganisms at submicrometer spatial resolution. The low throughput of this high-end topochemical analysis technique constrained the number of collected images for soil-incubated films to two images for each of the three PBAT variants including replicate films. We note that we did not exclude any data or outliers from our analysis.

Polyesters, monomers, soil, and enzymes

Polyesters were provided by BASF SE and synthesized as previously described (23, 24). The physicochemical properties of the polyesters are listed in table S2. To obtain similar ¹³C contents for the three PBAT variants (that is, PB*AT, P*BAT, and PBA*T), synthesis of all variants was performed with defined ratios of labeled to unlabeled monomers. The three PBAT variants were free of chemical additives.

The ¹³C-labeled monomers 1,6-¹³C₂-adipate and ¹³C₄-butanediol used for PBAT synthesis and for soil incubation studies were purchased from Sigma, with more than 99% of the indicated positions in the monomer containing ¹³C. We obtained 1-¹³C₁-terephthalate from dimethyl 1-¹³C-terephthalate purchased from Sigma. To obtain the free diacid, we dissolved dimethyl 1-¹³C-terephthalate in 2:1 water/tetrahydrofuran (5 mg in 2.4 ml), added 25 μl of a sodium hydroxide solution [37% (w/w)], and stirred the solution at room temperature for 2 hours. The solvent was then carefully removed under reduced pressure to obtain the hydrolysis product 1-¹³C₁-terephthalate (confirmed by ¹H NMR).

For PBAT and monomer incubations in soils under controlled laboratory conditions, we used agricultural soils from the agricultural center Limburgerhof (Rhineland-Palatinate, Germany). Physicochemical properties of the soils are provided in table S1. The soils were air-dried to a humidity of 12% of the maximum water-holding capacity of the soil, passed through a 2-mm sieve, and stored in the dark at 4°C for 12 months before use in the incubation experiments.

R. oryzae lipase was purchased as a powder from Sigma (catalog no. 80612). FsC was obtained as a solution from ChiralVision B.V. (Novozym 51032). Stock solutions of both enzymes in water were stored at −20°C.

Preparation of PBAT films and soils for incubation experiments

We prepared two sets of solvent-cast PBAT films that differed in the way that the PBAT films were attached to the silicon wafer substrates. For the first set, we solvent-cast PBAT films by adding three times 15 μl of a PBAT solution in chloroform [concentration, 5% (w/w)] onto a square-cut antimony-doped silicon wafer platelet (7.1 mm × 7.1 mm × 0.75 mm, Active Business Company). In between the additions of the polymer solutions, we allow the chloroform to evaporate. This procedure resulted in a PBAT mass of approximately 3 mg per wafer. Before incubation in soil, the solvent-cast polyester films were stored in the dark at room temperature for 1 week to ensure complete evaporation of the solvent (chloroform). PBAT variants from this first set were used for PBAT mineralization experiments (Fig. 2B), SEM imaging (Fig. 2C), and NanoSIMS imaging (Figs. 3 and 4 and fig. S8).

For the second set of PBAT films, we pretreated the silicon wafer platelets with Vectabond (Vector Laboratories, catalog no. SP-1800) before solvent casting of the polyester films. This second set of PBAT films was included to test whether the adhesion of the PBAT to the Si surface can be improved by this modified protocol. For the pretreatment, we exposed the wafers to a 1:50 diluted solution of Vectabond in acetone, subsequently dipped them into MilliQ water (Barnstead Nanopure Diamond), and dried them in a stream of N₂. PBAT variants from this set were used only to determine PBAT mineralization (fig. S1), but not for SEM and NanoSIMS imaging.

We prepared the soil for PBAT incubations by adding MilliQ water to the soil to adjust its water content to 47% of its maximum water-holding capacity. We subsequently transferred 60 g of the soil into each of the incubation vessels (100-ml glass Schott bottles). We prepared a total of nine incubation bottles in three sets of three bottles (see below). The soils were then preincubated at 25°C in the dark for 1 week.

After soil preincubation, we transferred the wafers carrying the solvent-cast polyester films into the soils in the incubation bottles. We added seven wafers to each incubation bottle. The wafers were spaced apart by at least 1 cm. All wafers were positioned upright in the soil. The three bottles of the first set each contained films of one of the three differently labeled PBAT variants obtained by direct solvent casting. The three bottles of the second set were identical to the first set except for the wafers, which were pretreated with Vectabond before solvent casting. The three bottles in the third set served as controls and contained soil but no PBAT films. All bottles were incubated for 6 weeks at 25°C in the dark. We note that our study therefore does not address potential effects of ultraviolet irradiation–induced changes in the structure of PBAT on its biodegradability. Over the course of the incubation, we gravimetrically determined the water content of the soils at defined time intervals. To sustain a constant soil water content, amounts of water that were lost from the soil through evaporation were replenished by adding corresponding amounts of MilliQ water.

Preparation and SEM imaging of soil-incubated PBAT films

After 6 weeks of incubation in soil, we carefully removed the silicon wafers carrying the PBAT films from the soils. To chemically fix the cells attached to the surfaces of the PBAT films, we directly transferred the films into a freshly prepared fixation solution (pH 7.4) containing glutaraldehyde (2.5%), sodium cacodylate (0.1 M), sodium chloride (0.1 M), potassium chloride (3 mM), and sodium phosphate (0.1 M). The films were exposed to this solution for 20 min at 25°C and subsequently transferred to a solution of OsO₄ in MilliQ water (1%) for 30 min of exposure on ice. Finally, we dehydrated the films in a series of water/ethanol solutions of increasing concentrations (70%, 5 min; 95%, 15 min; 100%, 2 × 20 min), followed by critical point drying of the samples with liquid CO₂ (Baltec CPD 030). Critical point drying resulted in detachment of the PBAT films from the wafer. To reattach the films to the wafers for further analyses, we used a double-sided adhesive, electrically conducting carbon tape (Ted Pella, product no. 16084-1). Directly after mounting the films onto the wafers with carbon tape, thin films of platinum (thickness, 10 nm) were deposited on the samples using a sputter coater (Baltec SCD 500). SEM was conducted on a Zeiss Supra 50 VP. Imaging was performed with a secondary electron detector at a working distance of 4.0 mm and an electron high tension of 5.0 kV. These films were also used for NanoSIMS analysis (see below).

PBAT films from the second set, for which wafers were pretreated with Vectabond before solvent casting of PBAT (see above), also detached from the wafers. We decided to reject further analysis of these films (that is, SEM and NanoSIMS).

PBAT film imaging by NanoSIMS

NanoSIMS measurements were performed on a NanoSIMS NS50L (Cameca) at the Large-Instrument Facility for Advanced Isotope Research (University of Vienna). Before data acquisition, analysis areas were presputtered by scanning of a high-intensity, slightly defocused Cs⁺ ion beam (beam current, 400 pA; spot size, approximately 2 μm). To avoid crater edge effects, scanning during presputtering was conducted over square-sized areas with an edge length exceeding the frame size of the subsequently recorded images by at least 15 μm. Every data set acquired on the soil-incubated polymer films contains image data recorded from (at least) two distinct depth levels, accessed by sequential presputtering with Cs⁺ ion fluences of 5.0 × 10¹⁶ and 2.0 × 10¹⁷ ions/cm², respectively. Application of the lower ion dose density enabled sampling of all cells within the analysis areas, irrespective of their size and/or morphology, whereas the extended presputtering allowed us to gain insight into cellular features contained within the lumen of bulky cells such as fungal hyphae (see, for example, Fig. 4).

Imaging was conducted by sequential scanning of a finely focused Cs⁺ primary ion beam (2-pA beam current) over areas ranging from 45 × 45 μm² to 70 × 70 μm² at a physical resolution of approximately 70 nm (that is, probe size) and an image resolution of 512 × 512 pixels. If not stated otherwise, images were acquired as multilayer stacks with a per-pixel dwell time of 1.5 ms per cycle. ¹²C⁻, ¹³C⁻, ¹²C¹²C⁻, ¹²C¹³C⁻, ¹²C¹⁴N⁻, ³¹P⁻, and ³²S⁻ secondary ions as well as secondary electrons were simultaneously detected, and the mass spectrometer was tuned for achieving a mass resolving power of >9.000 (according to Cameca’s definition) for detection of C₂⁻ and CN⁻ secondary ions. Image data were analyzed with the ImageJ plugin OpenMIMS, developed by the Center for NanoImaging (27). Secondary ion signal intensities were corrected for detector dead time (44 ns) and quasi-simultaneous arrival (QSA) of secondary ions. Both corrections were performed on a per-pixel basis. QSA sensitivity factors (“beta values”) were obtained from measurements on dried yeast cells, yielding 1.1, 1.06, and 1.05 for C⁻, C₂⁻, and CN⁻ secondary ions, respectively. Before stack accumulation, images were corrected for positional variations originating from primary ion beam and/or sample stage drift. ROIs were manually defined on the basis of ¹²C¹⁴N⁻ secondary ion signal intensity distribution images and cross-checked by the topographical/morphological appearance indicated in the simultaneously recorded secondary electron images (see fig. S10). While each cell from unicellular organisms was assigned to an individual ROI, image regions within the polyester surfaces and hyphae were segmented into multiple ROIs. Throughout the article, the carbon isotope composition is displayed as the ¹³C/(¹²C + ¹³C) isotope fraction, given in at%, calculated from the C⁻ and C₂⁻ secondary ion signal intensities via ¹³C⁻/(¹²C⁻ + ¹³C⁻) and ¹³C¹²C⁻/(2⋅¹²C¹²C⁻ + ¹³C¹²C⁻), respectively. Owing to superior counting statistics, all carbon isotope composition data shown in the article were inferred from C₂⁻signal intensities. We note that we did not observe any significant differences between ¹³C content values inferred from C₂⁻ signal intensities versus C⁻ signal intensities. For the line scan analyses displayed in Fig. 4, C₂⁻ normalized C¹⁴N⁻ signal intensities were used as an indicator of the relative nitrogen content {calculated via [¹²C¹⁴N⁻ (1 + ¹³C/¹²C)]/[¹²C¹³C⁻ + ¹²C₂⁻ (1 + (¹³C/¹²C)²)], whereby the term ¹³C/¹²C refers to the ¹³C-to-¹²C isotope ratio, calculated from the C₂⁻ signal intensities via ¹³C¹²C⁻/(2⋅¹²C¹²C⁻)}. This quantity formally refers to the relative nitrogen-to-carbon elemental ratio and was used in favor of the relative nitrogen concentration, which is inferable from C⁻ normalized C¹⁴N⁻ signal intensities, to minimize artifacts arising from the considerable topography within the areas of the fungal hyphae (28).

SUPPLEMENTARY MATERIALS

Supplementary material for this article is available at http://advances.sciencemag.org/cgi/content/full/4/7/eaas9024/DC1

Supplementary Materials and Methods

Fig. S1. Mineralization of PBAT films.

Fig. S2. NMR analysis of enzymatic hydrolysis products of PBAT films I.

Fig. S3. NMR spectra of terephthalate, adipate, and butanediol.

Fig. S4. NMR analysis of enzymatic hydrolysis products of PBAT films II.

Fig. S5. NMR analysis of enzymatic hydrolysis products of PBAT films III.

Fig. S6. NMR analysis of enzymatic hydrolysis products of PBAT films IV.

Fig. S7. Mineralization of terephthalate, adipate, and butanediol.

Fig. S8. NanoSIMS analysis of PBAT films after soil incubation I.

Fig. S9. Control experiment for NanoSIMS analysis I.

Fig. S10. Definition of ROIs.

Fig. S11. Control experiment for NanoSIMS analysis II.

Fig. S12. NanoSIMS analysis of PBAT films after soil incubation II.

Table S1. Soil characterization.

Table S2. Characterization of PBAT variants.

Supplementary Appendix. Calculations of the carryover during NanoSIMS measurements.

References (29–33)

This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial license, which permits use, distribution, and reproduction in any medium, so long as the resultant use is not for commercial advantage and provided the original work is properly cited.

Acknowledgments: We thank S. Probst, M. Jaggi, and F. Strasser for their help with growing E. coli, performing IRMS measurements, and NanoSIMS control sample preparation and data analysis, respectively. Funding: M.T.Z., T.F.N., R.B., H.-P.E.K., K.M., and M.S. thank the Joint Research Network on Advanced Materials and Systems of BASF SE and ETH Zürich for scientific and financial support. M.W. and A.S. were supported by the European Research Council Advanced Grant project NITRICARE 294343. D.W. was supported by the European Research Council Starting Grant project DormantMicrobes 636928. SEM imaging was performed at the Center for Microscopy, University of Zurich. Author contributions: M.T.Z., A.S., D.W., H.-P.E.K., K.M., and M.S. designed the study. M.T.Z., A.S., T.F.N., and R.B. performed experiments. All authors contributed to the writing of the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate our conclusions are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors.

• Copyright © 2018 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC).

 

 

 

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Is bamboo the answer to citrus greening in Florida ?

   Is bamboo the answer to citrus greening in Florida

At the present time, USA production of bamboo products provides but a tiny fraction of the national and international demand which is growing by geometric rates. By eliminating export requirements and import costs the domestic production of bamboo products is on a course to become a major force in Florida agriculture. Bamboo will become the next cash crop for Florida and beyond.  In a collaboration with an Italian bamboo company and Florikan, a producer of NASA chosen controlled release fertilizer we have been able to refine and accelerate  the production, propagation and growth of certain bamboo  species. Due to the elegance of design and safety and efficiency of this controlled release fertilizer it is adaptable to the growth characteristics of bamboo. Application is necessary only once or twice per year or per growth cycle. Edible bamboo shoots in large volume can be achieved in 3 years with mature trunks available for harvest in 5 years.  The future for Florida citrus farmers need not be bleak as they will be the logical recipients of this technology as the demand for bamboo products increases and as Florida farmers and growers realize the vast potential of this “perfect” plant. Just as the original settlers in Florida recognized a natural resource and improved upon it to create a major industry, the new pioneers are poised to begin a new frontier for Florida agriculture. 

Barrett Demler and James Demler Sweetgrass Farms, Sarasota, Florida, USA Published in Fruit Juice Focus Stefan@fruitjuicefocus.com 

Stefan Worsley e contact@juicemarket.info
t +44 (0) 7711 564 219

 

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Organic molecules on Mars

Organic molecules on Mars

On 6 August 2012, the Sample Analysis at Mars (SAM) instrument suite (1) arrived on Mars onboard the Curiosity rover. SAM’s main aim was to search for organic molecules on the martian surface. On page 1096 of this issue, Eigenbrode et al. (2) report SAM data that provide conclusive evidence for the presence of organic compounds—thiophenic, aromatic, and aliphatic compounds—in drill samples from Mars’ Gale crater. In a related paper on page 1093, Webster et al. (3) report a strong seasonal variation in atmospheric methane, the simplest organic molecule, in the martian atmosphere. Both these finding are breakthroughs in astrobiology.

Since NASA’s Curiosity rover landed on Mars in 2012, it has sifted samples of soil and ground-up rock for signs of organic molecules—the complex carbon chains that on Earth form the building blocks of life. Past detections have been so faint that they could be just contamination. Now, samples taken from two different drill sites on an ancient lakebed have yielded complex organic macromolecules that look strikingly similar to kerogen, the goopy fossilized building blocks of oil and gas on Earth. At a few dozen parts per million, the detected levels are 100 times higher than previous finds, but scientists still cannot say whether they have origins in biology or geology. The discovery positions scientists to begin searching for direct evidence of past life on Mars and bolsters the case for returning rock samples from the planet, an effort that begins with the Mars 2020 rover.

2018-07-25

NASA Statement on Possible Subsurface Lake near Martian South Pole

Science

Scientists have found evidence of a ‘lake’ on Mars

New evidence shows the presence of a large body of water, almost like a “lake,” one mile beneath the icy surface of Mars, according to a study published in the journal Science. They used data from Mars Express. If confirmed, this would be the largest body of liquid water found on Mars so far.Photo via @cnni

Far beneath the ice cap at Mars’s south pole lies a lake of liquid water—the first to be found on the Red Planet:

The view of Mars shown here was assembled from MOC daily global images obtained on May 12, 2003.

Credits: NASA/JPL/Malin Space Science Systems

Full image and caption

A new paper published in Science this week suggests that liquid water may be sitting under a layer of ice at Mars’ south pole.

The finding is based on data from the European Mars Express spacecraft, obtained by a radar instrument called MARSIS (Mars Advanced Radar for Subsurface and Ionosphere Sounding). The Italian Space Agency (ASI) led the development of the MARSIS radar. NASA provided half of the instrument, with management of the U.S. portion led by the agency’s Jet Propulsion Laboratory in Pasadena, California.

The paper, authored by the Italian MARSIS team, outlines how a “bright spot” was detected in radar signals about 1 mile (about 1.5 kilometers) below the surface of the ice cap in the Planum Australe region. This strong radar reflection was interpreted by the study’s authors as liquid water — one of the most important ingredients for life in the Universe.

“The bright spot seen in the MARSIS data is an unusual feature and extremely intriguing,” said Jim Green, NASA’s chief scientist. “It definitely warrants further study. Additional lines of evidence should be pursued to test the interpretation.”

“We hope to use other instruments to study it further in the future,” Green added.

One of those instruments will be on Mars later this year. NASA’s InSight lander will include a heat probe that will burrow down as far as 16 feet (5 meters) below the Martian surface. The probe, built by the German Aerospace Center (DLR), will provide crucial data on how much heat escapes the planet and where liquid water could exist near its surface.

“Follow the Water” has been one of the major goals of NASA’s Mars program. Water is currently driving NASA’s exploration into the outer solar system, where ocean worlds — like Jupiter’s moon Europa and Saturn’s moon Enceladus — hold the potential to support life. Even protoplanets like Ceres may explain how water is stored in rocky “buckets” that transport water across the solar system.

 Source : Inge Loes ten Kate Department of Earth Sciences, Utrecht University, Utrecht, Netherlands: i.l.tenkate@uu.nl in Science  08 Jun 2018: Vol. 360, Issue 6393, pp. 1068-1069

DOI: 10.1126/ science.aat2662

https://doi.org/10.1126/science.aat2662

Published By:

American Association for the Advancement of Science

Copyright & Usage:

Copyright © 2018, American Association for the Advancement of Science http://www.sciencemag.org/about/science-licenses-journal-article-reuse

This is an article distributed under the terms of the Science Journals Default License.

 

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