2024 Hydrological, chemical and biological assessment of two New Mexico headwater streams

Headwater streams play an important role in arid and semiarid regions. They provide freshwater to adjacent lowlands and temporarily store water as snowpack and groundwater. Downstream users – both humans specifically and ecosystems more broadly – depend on the delayed release, particularly during dry seasons. Headwater streams are highly vulnerable to climate change through shifts in snowmelt dynamics, changes in precipitation, evapotranspiration, and wildfire prevalence, among other factors. Monitoring headwater streams in dryland areas over time is therefore of critical importance.

Each year, graduate students enrolled in the Water Resources Program at the University of New Mexico monitor hydrological, chemical, and biological characteristics of two headwater streams in central New Mexico as part of a field methods class. This dataset contains measurements from the 2024 field campaign. At each stream site, measurements were repeated for two or three separate transects.

Las Huertas Creek (LH), the first monitoring site, is the only perennial stream in the Sandia Mountains. The stream drains north from the northeastern slope of the Sandias towards the town of Placitas, running through a narrow and heavily forested canyon. Three transects range in elevation from 2190 m – 2320 m above sea level and were sampled on September 28, 2024. The second monitoring site is on the East Fork of the Jemez River (JR) within the Valles Caldera National Preserve. The stream runs through the montane grasslands of the caldera, a 20 km circular depression formed by a large volcanic eruption and subsequent land subsidence approximately 1.2 million years ago. Two transects are located at an elevation of approximately 2560 m above sea level and were sampled on October 19, 2024.

Methods
Discharge, sediment, and channel geometry
Discharge at each transect was estimated using two methods: (1) the velocity-area method, with velocity measured using a Marsh-McBirney Flo-Mate Model 2000 flow meter, and (2) the salt dilution (or dry, or slug injection) method after Hudson and Fraser (2005). Electrical conductivity (EC) for the salt dilution method was monitored in 10-second increments using a YSI Professional Plus multiparameter meter, and in 1-second increments using an Arduino-based EC probe and datalogger built as part of the class. 

Particle size distribution of the channel bed surface was characterized based on the pebble count method (Wolman, 1954) by randomly selecting 100 pebbles along a zig-zag pattern (Bevenger and King, 1995) and measuring them with a gravelometer. In addition to the pebble count, grab samples (approximately 500 g) of stream and bank sediment were collected with a trowel at each transect and subjected to sieve analysis based on ASTM D6913 (ASTM, 2009). Three soil samples were taken along the vegetation transect in the adjacent riparian area outside of the stream channel. The soil texture by feel method was used to determine USDA texture class. 

One channel cross-section at each transect was surveyed using a Topcon AT-B3A auto level and stadia rod. Bankfull stage was estimated based on indicators such as change in vegetation and slope according to Harrelson et al. (1994). Centered on each cross-section, a longitudinal channel profile (20-30 m long depending on site conditions) was established by surveying 7-9 points along the channel thalweg. Bankfull discharge was then estimated based on Manning’s equation.

Water chemistry
The following water chemistry parameters were measured at five points (right bank, mid-right, center, mid-left, left bank) along a transect and averaged: temperature (°C), specific conductivity (µS/cm), pH, total dissolved solids (g/L), salinity (ppt), and dissolved oxygen (mg/L and %). The parameters were measured with a YSI Professional Plus multiparameter meter. Turbidity (NTU) was also measured at the same five locations described above using a LaMotte 2020i portable turbidity meter and averaged.

Water samples were taken to measure alkalinity in a lab environment; one liter of unfiltered water was collected at one location per transect and placed in a HPDE collection bottle, which was stored under refrigeration until lab testing. Alkalinity was determined by using endpoint titration; the titrant was 0.02 N sulfuric acid, while indicator solutions used were phenolphthalein (for carbonate) and bromocresol green (for bicarbonate).

Additional water samples were collected to test the anion and cation concentrations in a lab environment. At one central point per transect, approximately 1L of water was collected, filtered through a 0.45 µm glass fiber filter, and stored in a 50 mL centrifuge tube. A few drops of dilute nitric acid were added to only the cation samples, while nothing was added to the anion samples. All samples were stored under refrigeration until lab analysis. Cation concentrations were analyzed using inductively coupled plasma-optical emission spectroscopy (ICP-OES) with a PerkinElmer Optima 5300DV. Anion concentrations were analyzed using ion chromatography (IC) with a Dionex 1100 IC. Concentrations were then adjusted by subtracting positive field blank (i.e., background) values obtained running the same analysis on samples of deionized water left open to the air during each collection period.

Vegetation, organic matter and benthic macroinvertebrates
Foliar cover, basal vegetation cover, and bare ground were measured using a line-point intercept method (Herrick et al. 2020). Along a 25m transect perpendicular to the stream channel, observations were taken every 0.5 m (50 points in total).   At each point, a survey flag was placed next to the meter tape and every plant type was noted as well as overhead riparian cover. Taxa composition was estimated from LPI data. Percent canopy closure was measured at each transect, using a concave spherical densiometer. Measurements were recorded in the four cardinal directions and averaged. 

Organic matter was qualitatively sampled from three points (right bank, center, left bank) at each transect. Substrate was collected with a trowel. In the lab, substrate samples were processed for Ash Free Dry Mass (AFDM) (Steinman et al., 2017). Substrate samples were dried for 24 hours at 60°C, weighed, ashed at 500°C for 2 hours in a muffle furnace and reweighed. The difference in weights was calculated as the % organic matter as AFDM.

Aquatic macroinvertebrates were collected from a known area (measured in m2) at each transect using a D-frame net. Samples were preserved in 70% ethanol and stored in whirlpak bags.  In the lab, invertebrates were separated from organic/inorganic matter using forceps and a dissecting microscope, enumerated, and identified to Order (in most cases), using taxonomic references including Voshell (2002). Macroinvertebrate densities were calculated from raw counts as individuals/m2.

References
ASTM, 2009. Standard test methods for particle-size distribution (gradation) of soils using sieve analysis. ASTM D6913-04(2009)e1. West Conshohocken, PA.

Bevenger, G.S. and R.M. King, 1995. A pebble count procedure for assessing watershed cumulative effects (Vol. 319). US Department of Agriculture, Forest Service, Rocky Mountain Forest and Range Experiment Station, Fort Collins, CO.

Harrelson, C.C; Rawlins, C.L., and Potyondy, J.P., 1994. Stream channel reference sites: an illustrated guide to field technique. Gen. Tech. Rep. RM-245. Fort Collins, CO: U.S. Department of Agriculture, Forest Service, Rocky Mountain Forest and Range Experiment Station. 61 pp.

Herrick, J. E., Van Zee, J. W., McCord, S. E., Courtright, E. M., Karl, J.W. and Burkett, L.M., 2020. Monitoring manual for grassland, shrubland, and savanna ecosystems Second Edition – Volume I: Core methods. Retrieved from Las Cruces, New Mexico, U.S. Department of Agriculture, ARS Jornada Experimental Range, 77 pp.

Hudson, R. and Fraser, J., 2005. The mass balance (or dry injection) method. Streamline Watershed Management Bulletin, 9(1), pp.6-12.

Steinman, A.D., Lamberti, G.A., Leavitt, P.R., and Uzarski, D.G., 2017. Biomass and pigments of benthic algae, pp. 223-241 in (Hauer, F.R. and G.A. Lamberti eds) Methods in Stream Ecology: Vol. 1 Ecosystem Structure, Third Edition. Academic Press, Cambridge, MA.

Voshell, J.R., 2002. A Guide to Common Freshwater Invertebrates of North America. University of Nebraska Press, Lincoln, NE. 422 pp.

Wolman, M.G., 1954. A method of sampling coarse river‐bed material. EOS, Transactions American Geophysical Union, 35(6), pp. 951-956.