Diluting a solution is pretty easy. Usually you are taking a stock solution and adding a buffer or other solution to it to get the required dilution. The math is rather simple here as well.
Where:
Initial concentration or molarity.
Initial volume.
Final concentration or molarity.
Final volume
If math isn’t your strong suit, you can always use an online calculator to find volume of solution you need for your dilution.
When doing anti-body dilutions for IHC tests, everything is measure in micro liters (μL) and a micro-pipette is used to dispense the required volumes. Some common mistakes can occur such as, leaving out the antibody and allowing it to reach room temperature or not stirring/mixing the antibody before doing the dilution. Sometimes the antibody can settle/precipitate, so mixing should also occur once diluted and before being placed on an instrument.
Physio-chemical water analysis can be done many different ways. Water can be collected and sent off to a laboratory, or you can collect the data with hand held probes. If you require a more in-depth analysis it’s always a good idea to have it sent to a lab that specializes in water quality analysis, but if you are only doing a comparative study, it doesn’t make sense to spend all the money getting a professional lab to analyse the water quality.
While I was attending Minnesota State University: Moorhead (MSUM), we used may hand held probes on our assignments and student projects. We did a study on crayfish looking for preferred substrate size, and while doing that we recorded some simple water measurements such as: temperature, turbidity, and dissolved oxygen.
When working the life history study of the Mohave Tui Chub, we took many more readings. Temperature was taken on several sites due to the varying depth of the water. We wanted to make sure that the temperature did not interfere with the interaction between the chub and the mosquito fish. We also sampled dissolved oxygen, salinity, pH, and turbidity. Most of the measurements were made using a handheld probe. Some of the probes were multi functional, but some were uni-taskers. We took recordings several times per week at multiple points in the lake and ponds.
We also collected water samples in glass vials and delivered them to a professional water analysis laboratory in Los Angeles.
I worked for two successive summers at the HaPET office in the Bismarck, ND USFWS building. We were concerned with areas in North Dakota and South Dakota that were west of the Missouri river, or west river.
This project started as an offshoot of the Partners for Fish and Wildlife (PFW) program. This program provided help for landowners who had a lack of water on their property. The USFWS helped them build dams in water basins for irrigation and livestock.
Each year, before we started the survey work, we would directly call each land owner to make sure they would still allow us on their property. According to the agreement the landowner signed, we were allowed unobstructed access to the wetlands that were created, but we didn’t want to ruffle any feathers. Most folks were quite happy to have our help and would invite me in for coffee so we could talk about their land.
Then it became time to pound the pavement and get to all the wetlands. The surveys usually didn’t take too much time, but some were very hard to access. We were given an ATV, binoculars, and a spotting scope. The best way to survey the waterfowl were when they were on the water and not moving a lot. So sneaking up on them, or scoping them from a distance were the best methods.
Typical wetland, with a lot of waterfowl on it
These were similar to the four-square-mile surveys. I really enjoyed these surveys. I had to travel to the southern most parts of South Dakota, part of Montana, to the northern most parts of North Dakota. Every day was different, and I really enjoyed that. Some times we would be laid up at a hotel during inclement weather, but typically we could work through most weather conditions.
Green-wing teal male in full breeding plumage
We went out once in the spring to each wetland, and then again towards the fall to do a brood survey. The brood survey is super hard because all the adults have lost their breeding plumage, and the young hatch year birds, looked very similar to each other. It was much easier to ID them if the adults were around.
Late season plumage, it starts to get ratty looking before they molt their feathersMe holding two ducklings
My primary function while participating in the life history study of the Mohave Tui Chub was zooplankton counting. This was the first year of the field project, with several more years of study scheduled. During this phase of the project, we were developing a deeper understanding of the environment the chub lived in. We also did extensive water sampling and created micro habitats called mescosms. If you would like to read the dessertation paper Sujan Henkanaththegedara wrote about the Mohave Tui Chub, please see it here.
Conochilidae family – Rotifers
We collected zooplankton utilizing the horizontal water sampler which allowed us to get a representative sample at various depths across the waters were were testing. Once the water sample was collected, the invertebrates were strained out and placed in a sugar-formalin solution. Sugar formalin was chosen due to it’s similar osmostic characteristics to the saline environment they were taken out of.
Zooplankton counting station. The pink vials contain sugar-formalin and lots of invertebrates
The samples were than aliquoted out and a several mL sample was dumped in a zooplankton counting wheel. The wheel consists of a non-continuous circular line inset in a clear plastic dish. It is then placed under a dissecting scope and you start counting and classifying the zooplankton as you go around the wheel. We did this four times for each sample.
Zooplankton counting wheel
When we found an interesting or unknown specimen, we would take it over to a compound microscope on a glass slide. This allowed for much greater magnification so we could further classify the zooplankton.
Chydoridae family
For the study, we were not interested in classifying individuals down to the species, but instead got them all down to the family. Most of the samples contained many rotifers, and even on occasion we found some of the aquatic invertebrates such as water mites and chironomidae spp.
Chrionomidae sppHydrachnidiae spp. – Water mite
These photographs were taken through a compound microscope on a glass slide. I love using the Image-Based Key to the Zooplankton of North America, and recommend it’s use if you ever need to get down to the family of zooplankton you’re looking at. After that, a comprehensive key can be used to get to the species, but you’ll want to have several individuals for dissection, and a really good microscope.
While doing belt transects, this skill was essential to the research project. Without knowing what all of the grasses and leafy plants were, how could you determine if it was a native plant or not?
During the first month of employment on this project, the members of the team all went to local herbariums and studied identification keys so we could accurately identify all the foliage we came across.
Learning this skills was the most rewarding experience I’ve had at a job. The task was daunting in the amount of things you had to know. When I was unsure of something, I would take a picture from every single angle I could, or even dig out the plant by the roots so I could later identify what it was. This lead to many plant pressings and a huge amount of pictures. I was even working on a photographic guide to the wild flowers and also a guide to ligules. Ligules are used to identify grasses before the seed head has had a chance to grow. It’s the only way to determine which species a grass is in the springtime, even for cool season grass.
I did this for 9 hours a day for 5 months and learned an immense amount of information. I wish I could have retained it all. Some random images from the collection are below.
Geum triflorum-Prairie SmokeBasal foliage of G. triflorumLithospermum canescens – Hoary PuccoonCynoglossum officinale – HoundstongueElaeagnus commutata – Silver BerryArtemisia frigida – Pasture SageCerastium arvense – Prairie ChickweedPanicum capillare – WitchgrassWitchgrass closeup of flowerWitchgrass ligulePanicum capillare – WitchgrassWitchgrass liguleUnidentified liguleUnidentified ligule with long sheathLong sheath and smooth ligule – unidentifiedSphaeralcea coccinea – Scarlet GlobemallowToxicodendron radicans – Poison IvyAnother shot of the very hair, hairy ligule and leaf of this unidentified grass sp
I wish I could identify these grass species where I just have ligule pictures, but it’s nearly impossible without seeing the actual plant. I didn’t take enough pictures to identify it that way.
Carex family sedgeCarex liguleCarex ligule from another angleAmorpha canescens – Leadplant inflorescenceAmorpha foliage characteristics, part of the Fabaceae family (characteristic compound leaf structure)Lytta nuttalli – Nuttal Blister BeetleCypripedium candidum – Lady Slipper FoliageCypripedium candidum – Rare Lady Slipper on the North Dakota PrairieAnother Lady Slipper photoGreat shot of a rare prairie lady slipperWheatgrass sp based on clasping auricle presence – possibly quackgrassGaillardia aristata – BlanketflowerOnosmodium molle – False GromwellUnidentified hairy ligule with hairy leaf marginsAnemone canadensis – Anemone flowerAsclepias sp, probably incarnata – pink/swamp milkweedAsclepias spp. Probably sullivantii – Prairie MilkweedAsclepias speciosa – Showy MilkweedAsclepias speciosa – Showy MilkweedAsclepias spp. Probably sullivantii – Prairie MilkweedPenstemon digitalis or gracilis – BeardtongueHordeum jubatum – Foxtail BarleyLilium philadelphicum – Wood Lily, or at least a Lily speciesLily spp.Lactuca tatarica – Blue LettuceLactuca tatarica – Blue LettuceLactuca tatarica – Blue LettuceRosa arkansana – Wild Prairie RoseMonarda fistulosa – Wild bergamotRosa arkansana – Wild Prairie RoseBouteloua curtipendula – Sideoats GramaBouteloua curtipendula – Sideoats GramaBouteloua curtipendula – Sideoats GramaBouteloua gracilis – Blue GramaBouteloua gracilis – Blue GramaBouteloua gracilis – Blue GramaRatibida columnifera – Prairie coneflowerEchinacea angustifolia – Purple Coneflower – EchinaceaAndropogon geradii – Big BluestemAndropogon geradii – Big Bluestem – Turkey foot looking inforescenceAndropogon geradii – Big Bluestem liguleAndropogon geradii – Big Bluestem liguleSchizachyrium scoparium – little bluestemSchizachyrium scoparium – little bluestemCactus spp on the prairie
This was just a short list of the plants I saw and categorized. It’s unfortunate that I didn’t classify these plants shortly after I took the pictures. The grasses are hard to identify without having the whole plant in front of me, but during the survey period, I knew them perfectly. I wish I would have continued to create a ligule guide with key to the North Dakota prairie. The prairie there is a mix of short and tall grass, with hundreds of native grass species present, with nearly all of them being bunch grasses. Maybe I will get the opportunity again to classify the grasses and forbs of ND.
One of the best pieces of survey equipment I’ve used is the seine. I enjoy it so much because even the most dull stream is full of aquatic invertebrates and small fishes. The piece of equipment is composed of two long poles with a net strung between them. On the top side of the net there were floats, and on the bottom were weights.
Seining at the Buffalo River Science Center for a research project at MSUM
When I was working on creating a photographic guide to the fishes of the Buffalo River, a dear professor and I went out and seined many streams and ponds in the Buffalo River basin. If you have never tried it, it’s a great way to sample aquatic systems. It pulls up all of the macro invertebrates and the smallest fishes.
One of the first jobs I had when I was still in school was working for the Agricultural Research Service (ARS) at the Northern Great Plains Research Laboratory in Mandan, ND. They were a USDA affiliated lab, and I worked with a soil scientist on a crop rotation program. The project had gone on for quite some time, and I spent most of my days there on test plots making sure everything was going smoothly.
We started in spring by drilling in seeds, created quadrants, fertilized and tended those quadrants, and then harvested them in the fall. Since everything was laid out in a grid and we had information going back for years on everything including: soil moisture, nutrient/fertilizer loads, plant seed chemical analysis, and pesticide and herbicide use. This information was essential in creating a guide to which crops to plant for successive years. The office even produced a computer program for farmers to plan their fields out for years in advance.
After each quadrant was harvested, we used a shaker machine to separate the chaff, and then a laser seed counter to count how many seeds were produced. Those seeds then went to a grinder and then shipped out for analysis. It was so long ago that I did this work, Don Tanaka or Justin Hartel may be a better resource for questions.
One of my primary functions was bagging, counting, and organizing the seeds after they came off the sieve/shaker machine. The laser counters had to be calibrated and tweaked to get the most accurate count. So I spent quite a bit of time counting out 100 seeds and then running them through the machine to see how accurate we could get it. Weight wasn’t a reliable number due to each plant having different sized seeds.
When I was working on the native prairie project determining the percentage of invasive species present on WPAs, WMAs, refuge, and wilderness areas in ND, I had the opportunity to cover a vast swatch of prairie and catalog some of the species of gramminoids and forms that grew there.
I didn’t want to disturb some of the more rare species that might not have already been included in herbariums, so I took pictures of a lot of them. For some of the more common plants I took a plant press with me and collected many specimens of common grasses and forbs. The press is a simple device where you smash the plant down between two pieces of acid-free or news paper and then cardboard. On both sides of the cardboard, a piece of rigid wood or wood scaffolding is then bound with straps or screws. As the plant dries in between the sheets, it retains it’s shape and can be glued onto a clean sheet with all the information on where it was collected. The American Museum of Natural History has a great article describing the plant press.
Since it wasn’t feasible to press all of the plants, I took pictures of some of them as a digital catalog. Please check out the images below, and the images in the identification post.
Data entry and data logging are an integral part of science, and with that need came some experience using some of the databasing software out there. Excel is great for small personal projects, or simple recording of only a few categories, but sometimes you need more. For the waterfowl survey project I did with the USFWS, we used Microsoft Access to create forms that looked similar to our recording sheets to enter data into a table. Because the forms look identical, or nearly identical, to the forms we filled out in the field, it would be hard for someone to muck up the data entry by putting in the wrong column, which is a possibility within Excel. Having neatly organized data allowed us to run statistics to see if we can notice any trends going on.
Using access isn’t terribly hard if you have some experience using Excel. The form creation tool allowed me to easily create new forms for entering further information. Here is a video that shows you some very basic Access features.
Sharepoint also uses Access databases and is very powerful because it can easily be integrated at an organization that requires multiple people using the database concurrently. It plays very nicely with microsoft windows, but not sure about other devices. While I worked with Caris Lifesciences, we utilized the sharepoint service for almost everything we did outside of the Laboratory Information System (LIS). It was quite easy to create new forms and new logs to help us record anything from maintenance to reagent dilutions. If we ran into problems, or weren’t recording enough data, it was quite easy to add more fields to the forms without having to go back an amend all the previously collected information.
Part of the graduation program at MSUM required that we complete a capstone research program. It’s basically an undergrad thesis. Since I was graduating with an Evolution and Ecology degree, I wanted to conduct my own research on animal behavior. The test organism was Anax junius and the study criteria was conspecific cue and their reaction to it.
Green Darner dragonfly larvae
There was ongoing research going on between MSUM and another university to show that conspecific cue affected the behavior of fish. MSUM researchers found that adding mashed fish of the same species to the water induced fewer movements when food was present, which indicates the fish may be reacting to the cue in the water that a predation event had occurred nearby and that they should seek avoidance tactics to stay out of harms way.
I was intrigued by this research, so I decided I would do the same style of experiment on Anax junis, the green darner dragonfly. Aquatic systems are easy to use this cue in, so I collected many wild a. junius larvae and put them in plastic tubs with filtered water. A grid with small, but visible holes, was placed underneath each tub so we could record if a movement or feeding strike occurred.
The experiment started by introducing mashed up a. junius bodies via a syringe, and then worms were placed nearby the larvae and all movement was recorded for a set amount of time. I was able to get lots of student volunteers to help me collect data, so I had a good data set to start running some numbers on.
Experimental design
Here are the documents and presentations associated with this study. Please feel free to read them.
Initial concentration or molarity.
Initial volume.
Final concentration or molarity.
Final volume
