Patent Application
20190368978
This invention is not the subject of federally sponsored research or development.
Portions of the disclosure of this document contain material that is subject to copyright protection. The copyright owner has no objection to any reproduction of the document or disclosure as it appears in official records, but reserves all remaining rights under copyright.
A significant fraction of the microbial biosphere exists at elevated hydrostatic pressure (up to 110 MPa; 1086 atm) and low temperatures, yet an understanding of microbial adaptation to these conditions, particularly pressure, is limited (e.g., Wang et al., 2008). Though recent advances in molecular biology is beginning to address the problem it remains unclear whether adaptation to the piezosphere results from changes in a few genes, broader modification of the genome or effected mainly via regulatory processes (e.g., Simonato et al., 2006). What is required to become a piezophile is an interesting unanswered question. Most pure culture piezophiles under study have been subjected to decompression at some time during their isolation, based on studies by Yayanos & Dietz (1983), suggesting that piezophiles still possessing some ability to grow at normal atmospheric pressure would survive decompression and with obligate piezophiles, the quasi-first order lethal effects due to decompression is slow enough that isolates can be obtained if decompression times are minimized (˜90% loss of viability of an obligate piezophile when decompressed for 5 hr). Jannasch & Wirsen (1984) obtained non-obligate piezophiles from the deep oceanic water column in the absence of decompression (Jannasch, et al., 1982). Later deep-sea microbial ecology studies began to show large discrepancies between deep sediments 16S rRNA clone libraries and isolates obtained in the laboratory from the same source (e.g., Parkes, et al., 2009; Frye et al., 2008). On the other hand, members of enrichment cultures obtained from undecompressed deep-sea surficial sediment inocula and cultured at high pressure for several enrichment cycles did correspond to the gene library generated from the source sediment; identical inocula cultivated at sea surface pressures did not (Yanagibayashi et al., 1999). From the perspective of obtaining bacterial clones that are at all representative of the organisms residing in deep sediments it is becoming clear that procurement and culture must be effected at the pressures of the environment from where they came.
From a biogeochemical standpoint, hydrostatic pressure can also dramatically influence chemical gradients within microbial ecosystems, particularly in environments where metabolic, geothermal or hydrocarbon seep mechanisms result in elevated gaseous inputs (e.g., carbon dioxide, methane, other hydrocarbon gases, hydrogen sulfide) are driven into solution by pressure. Preservation of sediment samples from the deep oceanic seep environments is a particular challenge in that the time between sampling and retrieval can be hours and changes in pressure, temperature can result in substantial out-gassing that destroys the structural integrity of the retrieved sediment sample as well as changes the composition and activity of the contained microbial communities.
Technology developed in the laboratory of R. Sheryll, the Deep Ocean Benthic Sampler (DOBS) possesses a capability that is unique to the fields of deep-sea microbial ecology, biogeochemistry, and natural products biotechnology, the ability to obtain a contamination-free sediment core and preserve in situ conditions of pressure and temperature upon retrieval to the ship. By application of proposed mechanisms for obtaining multiple sub-cores at various depth horizons within the retrieved core samples, in the absence of decompression, permits in concert a) accurate assessment of the gaseous (e.g., hydrocarbons) and chemical (e.g., bicarbonate, hydrogen sulfide, etc.) gradients within the core without being disturbed by the “homogenizing” out-gassing that typically occurs in such samples when collected by conventional coring operations, b) the phylogenetic (DNA, ribosomal RNA, [rRNA]), functional (messenger RNA, [mRNA]) molecular study and culture of the resident microbiota using high pressure hardware available within the Woods Hole Oceanographic Institution (WHOI) and c) procurement of sediment samples for subsequent isolation of pure clones in the absence of decompression.
Deep Ocean Sampling: A historical background of approaches, instrumentation and rationale for the development of DOBS. For sampling in deep waters where hydrostatic pressure is a parameter that can affect microbial viability and growth, pressure-retaining samplers have been implemented in a variety of designs for retrieval of samples in the absence of decompression. In 1968 the first prototype high pressure water sampler was tested (Gundersen & Mountain 1972), followed by pressure-retaining samplers “with” (Jannasch et al., 1973; Jannasch & Wirsen, 1977) and “without” (Tabor et al., 1981) sample inlet protection to reduce the potential for contamination. The Jannasch & Wirsen (1977) pressure-retaining samplers retrieve concentrated (3 liters of seawater filter concentrated to 13 ml) undecompressed microbial samples from depths down to 6000 m. Bianchi et al. (1999) expanded the concept of Jannasch and co-workers to a multi-chamber setup that allow up to 8 pressurized samples to be taken during a single deployment, though inlet protection is less rigorous (alcohol-sterilized parafilm).
In 1978, Art Yayanos used a pressure compensating sample chamber to capture for the first time live deep-sea macrofauna (Yayanos, 1978). These deep-sea macrofauna were adapted for high pressure and low temperature, where limited food is available. These conditions were considered by Wirsen & Jannasch (1975) while working with psychrophilic bacteria at elevated hydrostatic pressures. Taylor (1979) found that culturing marine organisms under a high pressure oxy-helium atmosphere (50.7 Mpa; 500 atm) did not alter microbe viability, which led to the first hyperbaric isolation chamber (Jannasch, et al., 1982; Taylor, 1987) that allows, in the total absence of decompression, water column microbes collected from the deep sea to be isolated into pure clones using standard streak plate technique at pressures up to 60.8 MPa; 600 atm. Isolates are obtained from filter-concentrated deep sea water samples (concentrated 230×) or settled sediment slurry samples are streaked onto the surfaces of various solid nutrient media contained in multiple culture dishes mounted on a conveyer belt within the pressure housing. Given the ubiquity of hydrothermal vent and cold water seep discoveries in recent years there has been renewed interest in fluid sampling systems for procuring vent and seep samples for chemical analysis, including gas-tight isobaric sampling of hydrothermal fluids (Seewald and Doherty, 2001). There continues to be interest in systems for the collection and cultivation of deep-sea microbes in absence of decompression (Malahoff et al., 2002; Kato, 2006).
The importance of uncontaminated microbial samples for genomic research has led to the development of the Autonomous Microbial Sampler (AMS). This sampling system will obtain uncontaminated and exogenous DNA-free microbial samples from marine, fresh water and hydrothermal ecosystems (Taylor et al., 2006). The device is capable of obtaining six uncontaminated and exogenous nucleic acid-free, potentially hot (up to 350° C.) aqueous samples from most freshwater, marine and hydrothermal ecosystems. Samples are obtained via a heat-exchanging titanium nozzle possessing six sterile, nucleic acid free inlets that are protected by removable caps. During sampling the protective cap of a given inlet tube is hydraulically expelled via a sterile, nucleic acid-free aqueous snubbing fluid and a filtered sample from the environment subsequently obtained on an in-line filter. Complete protection of samples from exogenous contamination was demonstrated by passage of the sampling nozzle through seawater containing 106 cells ml−1 of a pigmented tracer organism into tracer organism-free seawater. While the AMS is able to protect samples from damaging temperature change it does not attempt to maintain environmental pressures upon return of the samples. AMS uses the method described by the Sheryll patent to obtain uncontaminated water samples.
Another apparatus, the Deep Sea Environmental Sample Processor (ESP), has been developed by MBARI (Roman et al., 2007). This micro-laboratory utilizes robotic technology and “standard tests to detect gene products and answer specific questions: By detecting specific RNA sequences, the system can find out which organisms are present. Or, by testing for particular proteins, it can ask what those organisms are doing. (For example, an RNA test can look for a particular kind of harmful algae; a protein test can learn how much toxin it is producing)” (http://www.mbari.org/mars/general/deep_esp.html).
The first pressurized deep oceanic sediment cores were obtained at the Blake Ridge and Carolina Rise 1995 during an Ocean Drilling Program (ODP) cruise (Leg 164) by use of the Pressure Core Sampler (PCS), a device that is capable of maintaining in situ pressures up to 681atm (69.0 Mpa; depth ˜6810 m) (Francis, 2001). In 2003 the Multi-Autoclave-Corer (MAC) and the Dynamic Autoclave Piston Corer (DAPC) were deployed in shallow water gas hydrate bearing sediments in the northern Gulf of Mexico. The devices retrieved, for the first time, near surface sediment cores under ambient pressure. The systems were designed to recover sediment cores at in situ pressures of up to 14.2 Mpa; 140 atm (MAC) and 30.4 Mpa; 300 atm (DAPC) (Heeschen et al., 2007). Parkes et al. (2009) developed a deep-isoBUG system used for culturing deep sub-seafloor sediments in absence of decompression (up to 25 MPa, 247 atm). Subsamples can subsequently be repressurized isolated and cultured at pressures up to 100 MPa (987 atm). Deep-isoBUG cooperatively works with the pressure retaining drill coring system called HYACINTH or PCS. The two systems are coupled together and a sub-core can be cut from the drill core while maintained inside the pressure retaining portion of the drill called the PRESS. The systems collectively allow sub-seafloor sediment samples to be sub-sampled and transferred into an isolation culture chamber of a design similar to the device described by Jannasch et al., (1982) or a chemostat for further processing and culture. Deep-isoBUG is essentially a subsampling transfer system. This system differs from DOBS in several ways; DOBS collects uncontaminated layer sediment core samples in absence of decompression for depths as great as 6,800 m, corresponding to a pressure 68.5MPa (>676 atm). Our Core Subsampling Unit (CSU, described later) works in a similar manner, where the subsample is transferred using a pushrod to move a subsample from one unit to the other. Surficial deep sea scoop sediment samples (˜5 ml) obtained by the manned submersible Shinkai 6500 have been successfully brought to the sea surface in a pressure and temperature retaining device (Kato, 2006) and enrichment cultures obtained in the absence of decompression (Yanagibayashi et al., 1999) in a large laboratory high pressure culture system called DeepBath (Kato, 2006). As indicated in the Introduction, several of the organisms found in the high pressure enrichments corresponded to the 16S rRNA clone library sequences found in the source sediments (not the case when the same sediments were enriched at 0.1 MPa, 1 atm); again relaying the importance of retaining in situ pressures in studies of the microbial ecology of the deep sea.
Generally lacking in deep-sea microbial ecology is a sampling approach that a) unequivocally guarantees procurement of uncontaminated sediment core samples from the deep sea, b) is able to maintain deep-sea conditions of low temperatures and high pressures upon return of the sample to the laboratory and c) possesses a mechanism for obtaining core sections under pressure to allow assessment of microbial diversity with depth in the sediment. To our knowledge, the only sampling technology in existence that possesses this unique combination of capabilities to the full depth of the ocean is the DOBS. Additionally, DOBS has been designed for flexible application; with some modification, the device can collect any type of sample found in the oceans whether it be live animals, minerals, water samples or sediment core samples, including the benthic boundary layer.
The invention provides improved elements and arrangements thereof, for the purposes described, which are inexpensive, dependable and effective in accomplishing intended purposes of the invention.
Other features and advantages of the invention will become apparent from the following description of the preferred embodiments, which refers to the accompanying drawings.
The invention is described in detail below with reference to the following figures, throughout which similar reference characters denote corresponding features consistently, wherein:
FIG. is a side elevational view of the embodiment of
This disclosure is not limited in application to the details of construction and the arrangement of components set forth or illustrated in the drawings herein. The disclosure is capable of other embodiments and of being practiced or of being carried out in various ways. Phraseology and terminology used herein is for description and should not be regarded as limiting. Uses of “including,” “comprising” or “having” and variations thereof herein are meant to encompass the items listed thereafter and equivalents thereof as well as additional items. Unless limited otherwise, “connected,” “coupled” and “mounted,” and variations thereof herein are used broadly and encompass direct and indirect connections, couplings, and mountings. “Connected” and “coupled” and variations thereof are not restricted to physical or mechanical or electrical connections or couplings. Furthermore, and as described in subsequent paragraphs, the specific mechanical or electrical configurations described or illustrated are intended to exemplify embodiments of the disclosure. However, alternative mechanical or electrical configurations are possible, which are considered to be within the teachings of the disclosure. Furthermore, unless otherwise indicated, “or” is to be considered inclusive.
General Description of DOBS: The Deep Ocean Benthic Sampler (DOBS;
The main mechanism by which DOBS can deliver sediment samples with their rRNA and short-lived mRNA signatures intact is by its ability to maintain the samples at in situ conditions during recovery to the sea surface and ship's laboratory. All of the physico-chemical conditions (e.g., temperature, pressure, chemical and redox gradients, etc.) that would alter microbe viability and the nucleic acid signatures, particularly that of mRNA, are maintained as though the sediment sample were still in situ. Physiologically and biochemically, the organisms are ‘unaware’ that they have been removed from the sea floor, at least for the trip to the ships laboratory. The DOBS achieves this primarily by maintenance of in situ pressure and temperature; the slowness of diffusion processes maintains the chemical properties of the sample. In situ pressure is maintained within DOBS by use of a helium gas (or other gas) cushion in a manner similar to how the Jannasch & Wirsen water samplers (Jannasch et al., 1973, Jannasch et al., 1976, Jannasch & Wirsen 1977, 1982) maintained deep-sea water samples at in situ pressures. Because gas is a compressible medium very little pressure change occurs within the sample during the inevitable dimensional changes in the vessel that occurs on its way to the sea surface.
DOBS consists of two separable pressure chambers (
Functioning of DOBS on the seafloor: A diagrammatic representation of DOBS functions are shown in
DOBS was built to maintain samples from depths of A800 m depth at in situ pressures, 66.4 Mpa (655 ATM) with a ≥4× safety factor. DOBS can safely hold gas pressures up to 2,620 ATM (266 Mpa) before elastic deformation occurs in the vessel. Additionally, DOBS is outfitted with a safety relief valve which will vent the contained gas long before these pressures would be reached (venting begins at user-determined pressures).
DOBS has been designed to accept additional SV's, allowing rapid recycling of the instrument and acquisition of additional cores while the core just retrieved is being processed. The SV can also be modified from its present design to contain multiple SAD probes for acquiring multiple core samples.
Technical Objectives DOBS System:
Construction of Core Subsampling Unit: Ultimately, a core subsample must be taken from the undecompressed core housed within the SAD probe at a given depth horizon for subsequent molecular or microbial analysis. Hardware for executing this operation is done by the Core Subsampling Unit (CSU) (or called the Sub Core Extractor (SCE)). The device, shown in
The following components of the invention shall be referred to by the immediately-following parenthetical abbreviation: Miniature Sub-Corer (MSC), Sample Vessel (SV), Acme Screw (AC), Sample Acquisition Device (SAD), Piston Core (PC), Nose Cone (NC), Core Tube (CT), Ball Valves (BV1), Deep Ocean Benthic Sampler (DOBS), Rotating Door (RD), Sample Module (SM), Ball Valves (BV), Unions (U), coupling magnets (DM), Core Liner (CL), Pressure Casing Wall (PCW), Magnetic Coupling (MC), Sample Tube (ST), High Pressure Tubing (HPT), Ledge to Anchor Piston (PA), Subsample Core (SC).
Prior to the sub-coring operation, the CSU and the SV of DOBS are disconnected as shown in
An advantage of the SM concept is that multiple units can be sterilized ahead of time and the entire core within the SAD sub-cored within a reasonable time without need for decompressing the CSU between sub-cores (there are two versions of the CSU or SCE one with pressure housing and one which does not require a pressure housing). The SM units can be quickly decompressed and sample core removed for subsequent molecular analysis. Two options are possible with this subsampling system: A), the sediment subsample contained within the SM is frozen while still pressurized by pouring liquid nitrogen over the tubing section. The sample can be stored in a pressurized, frozen state until processing. During sample processing the SM is quickly decompressed, the MSC removed and the frozen core ejected for further molecular processing. As an alternative, B), the MSC within the SM is not frozen, the unit decompressed, the MSC removed and the unfrozen core expelled and mixed with a chemical preservative (e.g., RNAlater; time required from decompression to preservation <15 sec).
A 3D representation of the sub-coring operation is diagrammatically illustrated in
Construction of the DOBS deployment platform: For acquisition of the sediment samples outlined above DOBS will need to be outfitted with a quadrilateral pyramid bottom lander similar to that illustrated in
To prevent cable entanglement with the DOBS during deployment syntactic foam cable floats (not shown in figure). These are attached to the ships hydro wire starting just above the attachment point of the sampler to prevent the cable falling onto the sampler when the hydro cable goes slack (see
Construction of adapters for interfacing the Core Subsampling Unit with high pressure culturing hardware within WHOI, especially the High Pressure Isolation Chamber (Jannasch et al., 1982; Taylor, 1987). For obtaining cultures from the DOBS samples, a hardware interface which allow the transfer of the core sample in the form of suspended slurry into the high pressure isolation chamber (Jannasch et al., 1983). See
The interface will be similar in design to the universal Sample Transfer System (STS) used by the Jannasch group to transfer water samples and cultures between all of the WHOI high pressure hardware (all have the same bolt pattern allowing transfer under high pressure between two available water samplers/culture chambers, the HPIC and a High Pressure Chemostat). The end caps of STS will be different from the Jannasch system allowing the mechanical transfer of the MSC (
Sub Core Extractor (SCE) Renamed Core Subsampling Unit (CSU)
Note: The some of the following components have had some of their names changed in later descriptions.
Originally I designed a pressure housing for a Sub Core Extractor (SCE) (renamed the Core Subsampling Unit (CSU) in later descriptions), which is used to remove pressurized sub samples from the Deep Ocean Benthic Sampler (DOBS). The internal configuration for the original design can be seen in
These components are held in place by the Forward End Plate Support (FEPS) and the Rear End Plate Support (REPS) which are separated by a series of four Linear Drive Columns (LDC). The forward and rear end plate supports are used to mount the gear motor and associated parts such as the Gear Shaft Support (SSG), drive gears, bearings, Limit Switches (LS) and the Acme Screw Bearing Covers (ASBC). A series of Push Rod Extractor Support's (PRES) are used to prevent the Push Rod (PR) from buckling due to forces that the Sub Core Probe Assembly (SCPA) are exposed to, when taking a pressurized sample from the DOBS.
This series of plates, Push Rod Extractor Support (PRES) follow the Push Rod (PR) as the Push Rod Block (PRB) moves forward or back keeping the push rod from buckling. The Push Rod Extractor Support's (PRES) collapse along with the Push Rod Block (PRB) as it moves forward keeping the Push Rod (PR) supported at all times. A cable is used (not shown) to pull the Push Rod Extractor Support (PRES) back as the Push Rod Block (PRB) is moved in the reverse direction, again keeping Push Rod (PR) supported to prevent buckling at all times.
The Push Rod Block (PRB) contains linear sleeve bearings which are fitted to the columns and are held in place by a Push Rod Bearing Covers (PRBC). The Push Rod Block (PRB) also supports a bronze nut which is attached to the Acme Screw (AS) used to drives the Push Rod Block (PRB) forward and back along the Acme Screw (AS). The Push Rod Block (PRB) is the terminal end support for the Push Rod (RP) as it is driven forward and back and moves the Sub Core Probe Assembly (SCPA) into the Sample Cavity (SC) of the DOBS where it extracts its sub-sample and returns this sub-sample to a pressure retaining sample chamber which is removable for later study.
As the push rod is advanced forward the Hard Stop Tracking Pin (HSTP) hits a hard stop inside the DOBS which pushes the piston to the back of the Piston Sub Sample Cavity (PSSC) (see
As the Push Rod (PR) moves forward (a) and the Hard Stop Tracking Pin (HSTP) hits a hard stop (not shown here, see
The Freeze Assembly Hard Stop (FAHS) shown in
At this point the two ball valves are closed sealing the forward portion of the Sub Core Piston Assembly (SCPA) with it sub-sample in between the two ball valves while maintaining the sub samples native pressure. This allows the removal of the Sub Core Piston Assembly (SCPA) from the high pressure conduit. Then the Ball Valve Freeze Assembly (BVFA) can be kept chilled and returned to the lab for further research. A new Ball Valve Freeze Assembly (BVFA) replaces the removed assembly for additional sampling.
The High Pressure-Sub Sample Storage Ball Valve (HP-SSSBV) and the High Pressure-Sub Sample Storage Location (HP-SSSL) along with the High Pressure Seal Cavity (HPSC) can be removed from the Sub Core Extractor (SCE). Where it can be maintained at temperature and pressure until the sample's return to the laboratory for transfer to the hyperbaric isolation culture chamber were additional culturing experiments can be carried out.
The high pressure sealing side of the Sub Core Extractor (SCE) or High Pressure Seal Cavity (HPSC) was designed to except two types of seals, one which requires that the Push Rod (PR) to have a very smooth surface finish and if blemished it could compromise the pressurized transfer of the subsample. As a backup, two interchangeable methods of sealing the high and lower pressure sides of the Sub Core Extractor (SCE) will be used. One method uses a spring energized seal; which will not leak unless the Push Rod (PR) is damaged. The other method uses a cup shape packing which will work even if the Push Rod (PR) is damaged. The draw back when using this type of seal is it may leak pressure wile sub-sampling. Having a backup method was implemented in case the Push Rod (PR) (used to penetrate the seals) is damaged in handling. If one of the Push Rods are damaged i.e., by scoring or scratching the outer surface of the rod. It can be replaced or the seal type can be changed to the cup shape seal which is not affected as much by scoring or scratching the surface of the Push Rod (PR). The Push Rod (PR) is coated using a diamond chromium coating to protect them from scoring or scratching.
In addition new methods will be attempted and implemented to save on the volume of gas used to charge the DOBS. Yielding a cost saving and reducing the number of gas cylinders normally required to charge the DOBS and the amount of time needed to fill the device. Reducing the volume of gas is also inherently safer.
One of these methods that will be attempted involves the replacement of the gas in the upper pressure chamber of the DOBS with a nonconductive silicone fluid. This can be done without changing the operation of the DOBS by simply changing the seals between the upper and lower pressure chambers of the DOBS with high pressure T shape bidirectional seals. This will allow the upper pressure housing of the DOBS to be pressurized to a greater pressure then the sampling depth, eliminating the need for pressure compensation of the fluid filled vessel. For depth greater than 5000 meters a compressibility studies will have to be conducted to determine if there is enough volume of gas in the lower pressure chamber of the DOBS to flush the snubbing fluid from the snubbing fluid chamber. This chamber is the contamination free interface between the inner and outer environments.
Use of the Core Subsampling Unit (CSU) was called the Sub Core Extractor (SCE): The basic concept behind the CSU is to procure “subsamples” at chosen depth horizons within the collected deep-sea core using a Miniature Piston Corer (MPC) that penetrates the core sample from the side (
As shown in
To take a core sub-sample from the Core Sample within the DOBS, a microprocessor controlled Motor & Gear Drive Assembly within the CSU is activated (
Once the core is solidly frozen the BVFA is decompressed, the MPC removed, and the frozen core ejected into a glass grinder tube (the surface of the MPC is quickly warmed to thaw a surface micro-layer of the frozen core so that it is releasable from the MPC). RNAlater is added to the grinder tube and the sub-core dispersed by the grinder tube pestle so that the instant the sediment sample is thawed the nucleic acids, including mRNA are instantly preserved, a critical capability for gene function studies made possible by the DOBS concept. The preserved sample is then extracted for DNA, RNA, mRNA according to the desired analyses to be conducted (i.e., for phylogenetic or gene functions studies).
The same sub-coring operation is followed for procurement of sample for subsequent biogeochemical analyses (e.g., CO2, CH4, H2S, etc.). If sensitivity permits, a fraction of the frozen core might be split, half for molecular analysis, half for biogeochemical analysis (e.g., GC-MS, etc.).
For culture work the freezing step is omitted. The sub-core can be decompressed and core contents dispensed into a dilution medium for resuspension and subsequent culture either at atmospheric pressure (1 ATM, 0.101 MPa) or in situ hydrostatic pressure in various culture media. It will also be possible to inject dilution medium into the BVFA to resuspend the sediment for subsequent transfer into a high pressure isolation chamber available at the WHOI laboratory should one wish to avoid decompression during the microbial culture/isolation process. The new method can be seen in
BVFA's are fabricated so that the DOBS Core Sample can be subsampled at multiple depth horizons (paired samples as close to one another as possible), providing a 5-point depth profile of the microbiology and biogeochemistry in the retrieved DOBS core (correspondence between molecular & biogeochemical analyses will be matched by interpolation).
Goniometer cart for the alignment of the CSU to the DOBS: The Goniometer Alignment Cart (GAC) (
Microprocessor Control of the Deep Ocean Benthic Sampler (DOBS):
A microprocessor (
DOBS Sub Sea Lander:
The DOBS lander (
The DOBS lander is deployed using two conductor torque balanced cable (diameter 1.73 cm, 0.68″) troll wire CC that could readily handle the combined weight of the DOBS (907 kg; ˜2000 lb.) and Lander (227 kg; ˜500 lb.). The cable was connected to the apex of the lander via a “Load Pin,” LP a strain gauge device that allowed real-time transmission of the weight of the assembly for detection of when the lander “touched down” on the sediment surface.
The lander (
Core Subsampling Unit (CSU) or Sub Core Extractor (SCE):
Sample Cavity:
Silicone Slip Cover also called Sub Core Tube Liner, Silicone Rubber Insert: shown in
Silicone Piston:
Miniature Piston Corer magnetic coupling:
Enhancement permitting larger diameter cores to be taken: The design of a 3.81 cm (1 W) diameter piston core tube option has been added for the study of gas hydrate and petrochemical sampling. DOBS sample probes can be exchanged while on the ship. This was previously not possible without the changes made to the Piston Corer Mechanism. This new enhancement will allow a wider and 5.1 cm (2″) longer cores [total elongated core tube length 15.24 cm (6″)] to be acquired.
Sample Acquisition Device (SAD probe) Without Piston Puller:
Sample Acquisition Device: The Sample Acquisition device (SAD) consists of a Sample Rod (SR) and a Sample Tube (ST) that slide out of a Sampler Housing (SH) during the acquiring of the sample and slides back into the Sampler Housing during the retrieving operation. For clarity, I have separated SAD in three parts for the purpose of explaining how this subassembly works, see
The relative rotation between the Sample Rod and the Sample Tube is being achieved by creating a channel, see Figure
Another feature of this type of mechanism, not described in this section, refers to a retractable key between the Sample Rod and the Sampler Housing. During the descend and the acquiring of the sample and also, a second retractable key between the Sample Rod and The Sample Tube that will become active after the Sample Cavity was closed. At this point both keys are envisioned as simple, one way, spring loaded retractable devices that will also allow manual reset of the SAD.
Quick Connect-Disconnect mechanism: The Quick Connect-Disconnect mechanism couples the Pushrod with the Sample Acquisition Device (SAD) during the descending of the Pushrod and also decouples the two parts after the SAD has been fully retracted into the Sampler Housing.
In
In
The Pushrod will decouple from the Sample Tube only when the Calibrated Steel Balls are free to move outward, pushed by the preloaded Plunger, and that position corresponds with the Sample Tube being in the initial position,
Description of Sample Acquisition Device (SAD): See Drawings 30A,30B, SAD 1, SAD 2. (The assembly is not limited to the method described here, other methods can be used to make this device as those skilled in the art of design).
The Sample Acquisition Device (SAD) collects an undisturbed sediments, power, liquid or surly sample. The Sample Tip/Door secures the Sample into the Sample Cavity and the Sampler is then retrieved back into the Sample Chamber. The figures SAD1, SAD2, show how this can be achieved by a simple push-pull action using a single linear actuator.
The Sampler Tube houses all the components of the Sample Acquisition Device (SAD) (see Drawings 30A, 30B, SAD 1, SAD 2). The Penetrating Tip (PT) is attached to the Sampler Rod (SR) at one end using a ⅛″ Dowell Pin (DP) held in place by two setscrews (SS). The Push Rod (PR) is mounted to the Sample Rod at its other end in the same manner (a ⅛″ Dowell Pin held in place by two setscrews). All the other components are mounted along the Sampler Rod between the Penetrating Tip and the Push Rod. The Sample Cavity (SC) is located inside a slot into the Sample Cam (SC) such that in the initial position, the actual Sample Cavity is aligned with the Sample Door (SD) of the Penetrating Tip.
The Sampler Retainer (SR) is keyed on top of the Sampler Cam. The Key Housing (KH) sits on top of the Sample Retainer such that the spring loaded Key (K) is keyed into the Sampler Tube (ST) slot and the spring loaded Plunger (P) is positioned 90° relative to the deepest point of the Sampler Retainer cam slot. In the initial position, the Plunger should be preloaded against the flat surface of the Sample Retainer.
The Key pivots about a 9/32″ Dowell Pin and is preloaded with a Torsional Spring (TS). There are two Teflon Washers (TW) added on both sides of the Key to reduce friction and prevent locking (not shown in the drawing). The Key is also free to rotate about its axis in counter-clockwise direction while it hard stops in the clockwise direction.
At the base of the Sampler Tube there is a Sampler Cam Key (SCK) screwed into the wall of the Sample Tube. The Sampler Cam Key is a commercial, stainless steel, spring loaded plunger. Its tip acts as a key and rides inside the Sampler Cam slot at all times.
The Push Rod has a threaded hole at its very end for hook up to a rod that will be pushed downward manually to simulate the action of a linear actuator upon the Sample Acquisition Device (SAD).
Sequence of Operation: SAD (
In the initial position, the Sample Acquisition Device sits above the sediment at a predetermined distance.
The operator (simulate the action of a linear actuator) pushes the Push Rod down. The Sampler should penetrate the sediment and continue it's descend with the Sample Cavity in open position until the Sample Cavity is filled with sediment. In parallel, the Sampler Cam Key is riding inside the straight section of the Sampler Cam slot. Next, the Sampler Cam Key engages the helical section of the Sampler Cam Slot. The Sampler Cam starts rotating counter-clockwise while the Push Rod, the Key Housing, the Sampler Rod and the Penetrating Tip are being held steady rotationally by the torsionally spring-loaded Key. This motion causes the closing of the Sample Cavity, which in turn traps the sediment sample inside the Sample Cavity. At the end of the rotational motion of the Sample Cam, the Plunger will pop-in inside the Sampler Retainer slot, locking rotationally the Sampler Cam with the Sampler Rod, Push Rod, Key Housing and the Penetrating Tip in the clockwise direction. When the Sampler Cam Key reaches the end of the Sampler Cam slot, the downward motion stops and the retrieving operation starts.
The operator will start pulling up on the rod coupled to the Push Rod. The Sampler Cam Key will force the Sampler Cam to rotate in the clockwise direction and along with it, the Push Rod, Penetrating Tip, Key Housing and Sampler Rod which are now coupled together in the clockwise direction by the Plunger.
The elements that were locked rotationally in the counter-clockwise direction during the downward motion are now allowed to rotate clockwise. The rotationally spring loaded Key will pivot counter-clockwise, rotate out of the linear slot and ride against the inside wall of the Sampler Tube.
The sequence is completed when the Sampler reaches the initial position. The only difference now is that the Sample Cavity is closed and the Sample Rod, Penetrating Tip, Push Rod and Key Housing are offset 90° rotationally.
To reset the mechanism, the operator must rotate the Push Rod in the clockwise direction 270° until the Key pops back into the Sampler Tube slot.
Not all mechanisms of the SAD probe can be seen from the drawings when viewed from in two dimensions. An example the Ramp of Plunger Ramp (PR) cannot be seen from the drawings.
The SAD Probe With Piston Puller Added: (See Attached Drawing Set
To take an undisturbed sample a piston #21 or 28 is used to reduce the motion of the sample as it moves vertically into the Sample Cavities/Sample Tube #11, 36. The Piston #21, 28 is not pulled as the name implies, instead it keeps the Piston #21, 28 stationary as the Sample Cam #5 moves through the Sampler Tube #1. This is accomplished by modifying the Sample Cam #5 and the Sampler Tube #1. The Sampler Cam #5 had an off set slot cut into the side of it. The length of the slot is based on the length of piston stroke. A longer the Sample Tube or Sample Cavity the longer the slot needs to be. The slot can expose the Sampler Rod #6 or 31 but it is not necessary for it to be exposed for proper operation. The Sampler Rod #6 or 31, if exposed can be used to center the Piston Puller #24, keeping it horizontal and tracking as it is displaced up and down through the slot. The Piston Puller #24 has two Pin Spring Plungers #13, inserted one on each side. These are used to control the start and stop of the piston travel (other controls can be used in place of the Pin Spring Plungers such as a solenoid or other method and it can be triggered from the outside of the Sampler Tube #5 the same for the location and type of the hard stops used i.e. other methods can be used in place of the Pin Spring Plungers). The piston puller has two rods #34 or rod/cables combination, screw, pined #25 or other method, into the bottom right and left side of its two lobes projected from the main body of the Piston Puller #24. These rods #34 are attached to the pistons #21 at the opposite end and as the Sample Cam #5 moves down into the medium to be sampled the pistons when triggered are displaced as the Piston Puller #24 moves in the slot. The triggers are set based on the depth of the sample to be taken and when to sample starts to enter the Sample Cavity #11, 36. They are controlled by the length of the helicoidal section of the Sample Channel in the Sample Cam #5 and a set of hard stops located on the inner wall of the Sample Tube (not shown) and the length of the slot. One or Two Ball Spring Plungers #17 located at the top and bottom of the slot in the Sampler Cam #5 which is used to pause the Piston Puller #24. When the hard stop is reached the Pin Spring Plungers #17 extends into the hard stop located in the inner wall of the Sampler Tube #1. This action locks the Piston Puller #24 inside the Sampler Tube #1 as the Sample Cam moves down inside the Sampler Tube #1. This action keeps the piston #21, 28 stationary as the Sample Cam #5 moves into the sediment. When the bottom of the helicoidal section of the Sample Tube Channel reaches the end of its helix this frees the Pin Spring Plungers #17 from their hard stops; as the end of the Sample Cam #5 rotation coincides with a ramp on the inner wall of the Sampler Tube #1 which allows the Pin Spring Plungers #17 to be freed allowing the piston to now move with the Sampler Cam #5 down into the sediment and retract back up into its starting position. The upper Pin Spring Plunger #17 then locks the Piston Puller #24 at the top of the Slot. If this upper lock false the sample can be lost depending on the type of material being sampled. So a Rod/Cable combination can be used. Having the lower portion of the Rod #34 replaced with a section of cable with a length equal to the length of the Sample Cavity that way if the lock false the Rod cannot push out the sample because the cable portion will not exert any force on to the Piston. A telescopic rod can be used in place of the Rod/Cable design.
This Piston Puller mechanism will work using a single or double piston design for that matter any number of pistons can used if the diameter allows for it. By exchanging different designed Sampler Cams different size samples can be taken. In addition by changing the design of the Sampler Rod #6 so as to except the larger Sample Cavity both types of Sample Cavities can be used. I made a Sampler Rod from #31, 35, from two sections in which I can replace one section for the larger Sample Cavity with another section that works with the duel Sample Cavity.
Compressibility Apparatus:
Compressibility Study of a Gas or Gas Mixture
The need to study organisms that exist in extreme environments as those found in deep-sea hydrothermal vents, cold water seeps and the abyssal plans are becoming increasingly important. Extremophiles are forms of bacteria, yeast fungi, Archaea and other organisms that live in these extreme environments.
These organisms are producing a variety of novel biopharmaceuticals, enzymes, small molecular weight and secondary metabolite chemical compounds. Along with their interesting DNA and genome information that are becoming increasingly beneficial to our industrial society.
Research into marine microbial ecology and the biodiversity of extreme environments is a promising area of study but is greatly restricted by the absence of affective sampling methodology. These restrictions include the ability to collect uncontaminated undisturbed samples and maintain the sample under ambient pressure and have the capability to manipulate this material in absence of decompression. In the design of the deep-sea sampling system, several unknown variables became a factor for the technology to become reality. First, the need to know specific molar volume of gases when compressed to extreme pressures (>600 atmospheres) and seconds, the gases used to create the internal atmospheric environments of the sampling system and its concentration.
These questions arise due to the manner of operation in which the sampling system retrieves uncontaminated undisturbed samples while maintaining the native environment of the sampled material.
To collect uncontaminated samples we must prevent the sampling probe from contacting the seawater during deployment and descent to the ocean bottom. This was accomplished by creating a controllable interface between the sampling chamber that contained a sterile sampling probe and the deep-sea outer environment. The interface consisted of a small-segmented tube connected together to the bottom entrance of the sample chamber. This tube is filled with a snubbing fluid (incompressible fluid), which is maintained in the tube by an end cap. This fluid is expelled upon reaching the desired sampling depth and replaced by a gaseous medium, which was contained inside the sample chamber during deployment and descent to the sampling location. To understand how these components work together to acquire an uncontaminated undisturbed sample a description of the operational characteristics of the sampling system and its method of operation was described above.
The Operational Characteristics of the Sampling System for the Extraction of Uncontaminated Samples:
The sampling system consists of three chambers that can be independently isolated from each other. They are connected and aligned vertically, one above the other. The top chamber is the motor housing which is in pneumatic communication with the sample chamber below its. This housing contains the drive mechanisms for opening the entrance of the sample chamber and deployment of the sample acquisition device (SAD). The sample chamber contains the SAD probe. This chamber can be removed from the other two chambers after a sample has been acquired for transport back to the laboratory for further study of the material inside. The third chamber or lid chamber is connected to the bottom entrance of the sample chamber and is in pneumatic and hydraulic communication with the sample chamber. It contains a series of cleaning devices used to wipe the SAD probe clean of debris as the probe is retracted back into the sample chamber after acquiring a sample. This lid chamber is the controllable interface between the entrance of the sample chamber and that of the deep-sea outer environment. It is filled with incompressible fluid called a snubbing fluid.
The DOBS uses gas to do work at the bottom of the ocean. When depths are greater than 600 atm the Idea Gas Laws no longer work to calculate the volume of gas needed to expel the snubbing fluid from the snubbing fluid chamber. There are no equations of state that work to calculate the super compressibility of the gas at that depth. The only way to figure out this problem is to use a device called the Barnett Apparatus or a better method, designed by me for this work called the Compressibility Apparatus see
The solution was to determine the compressibility factor by empirical methods. I designed a simple experiment that would determine the compressibility of any gas or gas mixture of interest. This can be done by taking a small length of high-pressure tubing say 15 cm with an inner diameter of from 0.3175 cm to 0.15875 cm. At each end attached are two-way valve the bottom connected to a high-pressure hand pump with an accurate pressure gauge and reservoir pressure cylinder. The valve on top in turn would be fed into an upside-down volumetric flask filled with distilled water. This flask would also have an exit to allow the displaced water to leave the flask. This flask would need to be about 800 times the volume of the piece of tubing used as the pressure vessel. The system could be heated or chilled to the temperature of interest. We fill the reservoir pressure cylinder with a gas or gases to be tested. Using hand pump we can compress the gas by hydraulically introducing glycerol into the reservoir pressure cylinder. Which will displace the test gas from the reservoir pressure cylinder into the piece of high-pressure tubing. When the desired pressure is reached as seen on a pressure gauge, the system is given time to reach a temperature equilibrium. Once occurred, the desired pressure and temperature has been reached, the bottom valve can be closed and the top valve can be opened slowly allowing the compressed gas to empty into the water filled flask as the water is displaced and the system is allowed to come to equilibrium the new decompressed volume of gas can be read directly from the volumetric cylinder.
We need to determine the total inner volume of our test vessel. To this volume we need to consider the stress and deformations in the pressure vessel. With this information we could calculate the actual compressibility factor for the gas being tested.
I have built this test device and used it to test the actual compressibility factor of the Oxy-helium mixture intended to be use.
Compressibility Apparatus
This picture shows the experimental apparatus for determining the compressibility of the Oxy-Helium gas mixture. Starting from the lower left-hand side of the picture showing: 1) Glycerol Reservoir (white bottle), above that; 2) hand operated high-pressure pump, above to the right; 3) Purge Valve V8, to the right and above; 4) Pressure Gauge, to the rights and down; 5) Valves V1, V2, moving downwards; 6) Reservoir Pressure Cylinder (RPC), moving right; 7) Valves V3 and Purge Valve V10, moving right; 8) Pressure Gauge, moving upwards; 9) Valves V4, V5, between V5 and below valve V6; 10) Experimental Pressure Test Cylinder (EPTC) above this is; 11) Valve V6, and directly above is valve V7 which is a metering valve, connect to V7 is, 12) Volumetric Flask for measuring volume of decompressed gas; 13) Water Bath which contains valves V4-V7. 15) A Recalculating chiller/heater not shown here is connected to the insulated water bath 13. On the top center is; 15) Purge Valve V9, below to left; 16) Inlet check valve to helium supply, to the right; 17) Oxy-Helium Inlet port.
Deep Ocean Benthic Sampler (DOBS) possesses a unique capability to the fields of deep sea microbial ecology and natural products biotechnology, the ability to obtain a contamination-free benthic boundary layer sediment core samples and preserve in situ conditions of pressure and temperature upon retrieval to the ship.
Broader Impacts
Availability of DOBS to Scientific Community. Biological diversity of marine organisms, especially of microbes found in the deep-sea, such as deep-sea hydrothermal vents, deep-sea cold seeps, but also the understudied abyssal plains holds great potential for natural products discovery and applied biotechnology. General cultivation independent diversity assessments based on the universal taxonomic marker molecule 16S rRNA have revealed a vast diversity of microorganisms. However, for the most part the physiology and metabolic abilities of these organisms remain unknown. This is mainly due to the fact that only a small fraction (<1%) of these microbes can be successfully cultivated in the laboratory, severely limiting our understanding of the ecological role and metabolic potential of these organisms. This is particularly relevant for the deep-sea, where organisms exist that are adapted to the in situ conditions, including high pressures and low temperatures. This prevents many, if not most microorganisms from these environments from being cultured in the laboratory, suggesting that we have only scratched the surface of the metabolic potential and the extent of physiological diversity of the microbial communities inhabiting these environments. Using its innovative design and methodology, DOBS will allow access to the microbes from these environments by being able to preserve in situ conditions, enabling their cultivation and thus the application of genomic and postgenomic techniques. These microbes may represent a rich reservoir of so far untapped biodiversity with obvious implications for bioprospecting. In addition, DOBS will provide extremely useful in sampling gas- and gas-hydrate bearing sediments. By maintaining in situ conditions the distribution and activity of microbes can be directly correlated with the concentration of volatile and unstable compounds, such as gas hydrates, providing novel insights into the microbial biogeochemistry of these habitats. Gaining a better understanding into the role that microbes play in affecting hydrate stability and dissolution is of great relevance. Finally, the technology behind DOBS will be made available to the scientific community by Cyclops Research and Development, Inc.
Introduction: Sampling Deep Sea Microbial Biosphere
A significant fraction of the microbial biosphere exists at elevated hydrostatic pressure (up to 110 Mpa; 1086 atm) and low temperatures, yet an understanding of microbial adaptation to these conditions, particularly pressure, is limited (e.g., Wang et. al., 2008). Though recent advances in molecular biology is beginning to address the problem it remains unclear whether adaptation to the piezosphere results from changes in a few genes, broader modification of the genome or effected mainly via regulatory processes (e.g., Simonato et. al., 2006). What is required to become a piezophile is an interesting unanswered question. Most pure clone piezophiles under study have been subjected to decompression at some time during their isolation, based on studies by Yayanos & Dietz, 1983, suggesting that piezophiles still possessing some ability to grow at normal atmospheric pressure would survive decompression and with obligate piezophiles, the quasi-first order lethal effects due to decompression is slow enough that isolates can be obtained if decompression times are minimized (˜90% loss of viability of an obligate piezophile when decompressed for 5 hr). Jannasch & Wirsen, 1984 obtained non-obligate piezophiles from the deep oceanic water column in the absence of decompression (Jannasch, et. al., 1982). Later deep-sea microbial ecology studies began to show large discrepancies between deep sediments 16S rRNA clone libraries and isolates obtained in the laboratory from the same source (e.g., Parkes, et. al., 2009; Frye et. al., 2008). On the other hand, members of enrichment cultures obtained from undecompressed deep-sea surficial sediment inocula and cultured at high pressure for several enrichment cycles did correspond to the gene library generated from the source sediment; identical inocula cultivated at sea surface pressures did not (Yanagibayashi et. al., 1999). From the perspective of obtaining bacterial clones that are at all representative of the organisms residing in deep sediments it is becoming clear that procurement and culture must be effected at the pressures
From a biogeochemical standpoint, hydrostatic pressure can also dramatically influence chemical gradients within microbial ecosystems, particularly in environments where metabolic, geothermal or hydrocarbon seep mechanisms result in elevated gaseous inputs (e.g., carbon dioxide, methane, other hydrocarbon gases, hydrogen sulfide) are driven into solution by pressure. Preservation of sediment samples from the deep oceanic seep environments is a particular challenge in that the time between sampling and retrieval can be hours & changes in pressure, temperature can result in substantial out gassing that destroys the structural and microbial integrity of the retrieved sediment sample.
Technology developed in the laboratory of R. Sheryll, the Deep Ocean Benthic Sampler (DOBS) possesses a capability that is unique to the fields of deep sea microbial ecology and natural products biotechnology, the ability to obtain a contamination-free core and preserve in situ conditions of pressure and temperature upon retrieval to the ship. By application of the Core Sub-Sampling Unit (CSU) mechanisms for obtaining multiple sub-cores at various depth horizons within the retrieved core samples, in the absence of decompression, permits in concert a) accurate assessment of the gaseous (e.g., hydrocarbons) & chemical (e.g., bicarbonate, hydrogen sulfide, etc.) gradients within the core without being disturbed by the “homogenizing” out-gassing that typically occurs in such samples when collected by conventional coring operations, b) the phylogenetic (DNA, ribosomal RNA, [rRNA]), functional (messenger RNA, [mRNA]) molecular study and culture of the resident microbiota using high pressure isolation culture chamber and chemostat available within the Woods Hole Oceanographic Institution (WHOI) and c) procurement of sediment samples for subsequent isolation of pure clones in the absence of decompression.
In the grant we proposed to develop DOBS into a routine instrument that can be used by the oceanographic community to obtain undisturbed sediment cores maintained under in situ conditions for biogeochemical and microbiological analyses. Specifically we will address the following objectives:
The invention is not limited to the particular embodiments described and depicted herein, rather only to the following claims. I claim:
This Application incorporates by reference and, under 35 U.S.C. § 119(e), claims priority to U.S. Provisional Patent Application Ser. No. 62/471,482 filed on Mar. 15, 2017. This Application incorporates by reference U.S. Pat. No. 5,559,295, issued Sep. 24, 1996.
