The hyporheic mesocosms facility (HMF) is located approximately 150 m downstream of the WS1 well network (see https://andlter.forestry.oregonstate.edu/data/abstract.aspx?dbcode=CF011). The HMF is located on a streambank and consists of twelve identical pipe segments. Each segment is a 1-m long hollow aluminum pipe with internal diameter of 20.32 cm. Pipe segments are capped with high density polyethylene (HDPE) end caps. A 0.5-cm diameter hole in the center of each end cap allowed flow into and out of each segment. The bottom and top caps were identical and served to spread the point source of water at the inlet into uniform laminar flow across the full width of the mesocosm and then collapse that flow back to the outlet point, thus limiting the development of preferential flow paths and large dead zones adjacent to the end caps. To accomplish this, 18 grooves, spaced 20° apart, radiated outward from the central hole, alternating in length from 3.8, 5.7, and 8.0 cm. Each groove was narrow and shallow at the inlet hole and gradually widened and deepened along its length. A diffuser plate was placed over the grooves in the end caps to keep them free of sediment. The 20.12-cm-diameter diffuser plate was made of sintered stainless steel with high-flow square weave support layers and a nominal pore diameter of 40 µm.
Each hyporheic mesocosm was packed with native streambed sediment in May 2016. We collected the sediment used to pack the mesocosm segments from the bedload trap basin located at the mouth of the WS01 catchment, ~50 m downstream from the WS1 stream gage. The basin sorts sediment, with the coarsest bedload being dropped at the head of the basin and the finest sediment and most organic matter accumulating in the deepest and most distal end of the basin. We chose a location about 1/3 of the distance along the length of the basin, where surface sediment was dominated by fine gravel, sand, and finer textured mineral sediment but with little to no obvious accumulation of organics. In August 2014, the bedload trap basin was drained and allowed to dry for several days. After this drying period, we dug moist sediment from the floor of the basin and sieved it through galvanized wire mesh with square openings measuring ~6 mm (1/4 inch) on a side to remove all large particles. Standing water was not present over the excavated material, so fines were not washed from the sediment as it was dug and sieved. After sieving, we transferred the sediment to woven polypropylene sandbags, which we then layered along the upstream face of the dam so that they would be underwater once the catch basin was refilled a few days later. The sediment was thus stored under water until we were ready to pack the mesocosms.
We packed the mesocosms in May 2016. We first retrieved sandbags from the pond and allowed them to drain by gravity. The woven polypropylene material is relatively tight, so the sandbags drained slowly with little loss of fine materials. To ensure homogeneity during packing, we emptied 2 to 3 sandbags into a plastic tub and mixed them with a shovel. Then we dumped a small scoop of sediment (~500 g) into each of the 12 mesocosm segments. A 2nd scoop was added to each of the 12 segments, and the mesocosms were ultimately filled by continually adding scoops in sequence. This sequential packing was intended to spread any variation in sediment texture or organic matter content evenly across all 12 pipe segments. Once the tub was empty, we used a long-handle square point tamper (10.16 cm x 10.16 cm) to compact the layer of sediment in each pipe. Then, we refilled the tub with sediment and repeated these steps until all 12 pipe segments were full. In total, 24 sandbags of sediment were needed to pack all 12 pipe segments.
The mesocosm segments were held vertically on an aluminum rack, and stream water was pumped to a head box >3 m above the mesocosms to provide constant head to drive flow upward through the mesocosms. Each mesocosm segment was connected with polyethylene tubing (internal diameter of 0.43 cm), first running from a main water supply pipe fed from the head box and into the bottom of the 1st mesocosm segment, and then from the top of the 1st segment to the bottom of the 2nd segment. Outflow from the top of the 2nd segment was regulated with a high precision needle valve (HOKE®, Spartanburg, South Carolina; part number 1335M4Y; Milli-mite 1300 Series valve with a globe flow pattern, in stainless steel, with a 1° stem and 0.047-inch orifice with Cv (flow coefficient) = 0.01, CRANE Instrumentation & Sampling PFT Corporation, Beijing, China). The total flow path from the inlet of the 1st mesocosm segment to the outlet of the 2nd segment defined a 2-m hyporheic flow path. We assumed that the tubing connecting the segments of each mesocosm had minimal influence on biogeochemical processing compared with the combined length of the 2 pipe segments because of the tubing’s limited surface area and short residence times.
We maintained flow rates through each mesocosm as close to 48 mL/min as was possible throughout the duration of the study (May 2016–Sept 2018). We measured flow velocities with tracer tests showing that median travel times through the 2-m mesocosms ranged from 9.12 to 13.87 h (mean = 10.43 h, s = 1.06 h) across all mesocosms on all sample dates (Table 2). Thus, with a flow rate of 48 mL/min, the mean flow velocity through the mesocosms was 0.19 m/h and ranged from a low of 0.18 m/h to a high of 0.21 m/h, which closely matched flow velocities observed in the well network during tracer tests.
The mesocosms were instrumented with a variety of in-line sensors to provide real time monitoring. The main water supply line was split into 3 sub-lines, each feeding a pair of mesocosms. Each of the 3 sub-lines included an in-line EC sensor (CS547A-L, Campbell Scientific®, Logan, Utah), a venturi mixer (6.4-mm Venturi Injector, A2Z Ozone®, Louisville, Kentucky), and an injection port. In-line EC sensors were also located at the outlet of all 6 mesocosms along with an electronic flow meter to monitor flow rates in real time. Sampling ports were located on the inlet tube to each individual mesocosm, between the 2 segments of the mesocosm, and at the outlet from the 2nd segment thus allowing us to sample at flow path distances of 0, 1, and 2 meters.
Sampling the mesocosms was a multistep process. We first made in situ measurements using sensors for temperature, dissolved oxygen, pH, and EC, after which we collected the actual water sample. To make our sensor measurements, we stopped flow downstream of the sample port by closing a valve, and we opened the sample port so that the sample collection rate was close to 48 mL/min—the same rate as the flow through each mesocosm to minimize the potential to develop preferential flow paths through the sediment when sampling. We measured dissolved O2 and temperature in an ~15-mL flow-through cell containing the probe end of a YSI ProODO Optical Dissolved Oxygen Meter (YSI® Incorporated). We measured pH and EC in ~20 mL of water collected in a graduated cylinder using either a YSI Model 60 pH (YSI® Incorporated) or a Hach® H160NP Portable pH meter, and EC with a ProfiLine Cond 3110 (WTW® Wissenschaftlich-Technische Irkstätten GmbH).
To collect the water sample, an acid-washed 60 ml BD® syringe was connected to the sampling port and water was collected roughly at 48 ml/min by manually pulling on the plunger of a sample syringe. Approximately 120 ml (two syringe full) of water was used to rinse sample syringe, filter apparatus, and ash-free GFF filter twice. Another 60 ml of water was collected and rinsed an acid-washed 250 ml HDPE Nalgene® bottle two times. Then 250 ml of filtered water was collected in the HDPE Nalgene® bottle and 60 ml unfiltered water was collected in sample syringe fitted with air tight 3-way luer lock stop valve. Note that we used the same technique to collect filtered and unfiltered stream water samples and we also collected field duplicates from both the stream and mesocosms for quality assurance. We stored samples in an ice chest kept cold with ice packs and then transported them to the lab where they were refrigerated at 4ºC until analyzed.
Before February of 2018, we sampled all 6 inlet ports, then the intermediate ports, and finally the outlet ports. Sampling required 0.5 to 1 h at each location (~3 h total sampling time), whereas the mean travel time through 2-m mesocosms was 10.43 h. We modified our sampling protocols to support characterization of changes in parcels of water moving through the mesocosms (i.e., Lagrangian sampling) starting in February 2018. First, we intentionally timed rounds of sampling to coincide with the travel time of water flowing through the mesocosms, waiting 5 to 6 h to sample the intermediate ports and another 5 to 6 h to sample the outlets. Second, to reduce the time needed to collect field samples, we designed a sampling system that consisted of 3 sets of 6 acid-washed sample bottles (500-mL HDPE Nalgene®; Nalge Nunc International Corp., Rochester, New York). We used each set of bottles to collect water from the 6 inlet, intermediate, and outlet ports. A set of 6 sample bottles would be connected to 6 sampling ports to collect ~500 mL of unfiltered water, regulating the flow rate to ~48 mL/min by using the valve on the sample port. The 500-mL sample bottles were rigged with inlet and outlet tubes that fit tightly into holes drilled into the bottle caps to minimize chances for contamination when collecting the water sample. The actual water sample was then collected using a 60-mL syringe connected to the outlet tube of the 500-mL bottle rather than directly to each mesocosm’s sampling port. See Serchan (2021) for additional details.
All laboratory analyses followed CCAL’s standard operating procedures and thus are consistent with other water chemistry data collected at the H. J. Andrews Experimental Forest (for example, dataset CF002 Stream chemistry concentrations; https://andlter.forestry.oregonstate.edu/data/abstractdetail.aspx?dbcode=CF002).
The CCAL standard operating procedure for DOC and DIC analyses were developed from American Public Health Association (APHA) methods. Citations for the methods used in DOC and DIC analyses will be in the following format: (CCAL standard operating procedure, APHA method, comparable EPA method, method detection limit).
Detailed descriptions of the methods and references to the procedures are given in Serchan (2021) and copied here:
Unfiltered syringe samples collected in the field were analyzed for DIC. Prior to DIC analysis, stopcocks were removed from syringes and immediately replaced with 25 mm diameter VWR® Syringe filters with polypropylene housing. Sample water in syringe were pushed through filter into an acid-washed 40 mL borosilicate vial. The vial was filled by holding it at an angle so sample water ran down its side wall. When vial was close to being full, it was straightened, filled to its brim, and capped as soon as sample formed inverted meniscus at its mouth. Filtered samples were then analyzed on a Shimadzu TOC-VSCH Combustion Carbon Analyzer within 72 hours (CCAL 21A.1, n/a, n/a, 0.05 mg C/L).
Field filtered water samples in 250 mL Nalgene® bottles were analyzed for DOC. An aliquot (~25 mL) of field-filtered 250 mL sample was analyzed for DOC. Aliquots were poured into baked 40 mL borosilicate vials and analyzed on a Shimadzu TOC-VSCH Combustion Carbon Analyzer (CCAL 20A.3, APHA 5310B, EPA 415.1, 0.05 mg C/L).
Citations for all Cooperative Chemical Analytical Lab (CCAL) procedures are shown at: https://ccal.oregonstate.edu/methodology
Johnson, S.; Fredriksen, R. 2019. Stream chemistry concentrations and fluxes using proportional sampling in the Andrews Experimental Forest, 1968 to present ver 23. Environmental Data Initiative. https://doi.org/10.6073/pasta/bb935444378d112d9189556fd22a441d.
Serchan, Satish P. 2021. Evidence of Buried Particulate Organic Carbon as Foundation for Heterotrophic Carbon Metabolism in the Hyporheic Zone of a Montane Headwater Stream in the H. J. Andrews Experimental Forest, Oregon, USA. Corvallis, Oregon: Oregon State University. 91 p. M.S. Thesis. (Pub No. 5369)
