Information

When is it unacceptable to measure dry mass in an experiment to measure growth of plants?


I'm an A level student (British high school final year) and I was doing an exam question which asked for the design of an experiment to test the effect of light intensity on the growth of a certain type of plant. Here is the paper; it is question 2(c). The marking scheme can be found here.

As my measured dependent variable, I picked dry mass. However, the marking scheme specifically says to 'ignore dry mass' and asks for use of 'mass'. We were always taught that dry mass is the most accurate measure for experiments where we are trying to measure the effect of a certain parameter on the growth of plant. So now I am wondering - when is it unacceptable to use dry mass as a measured dependent variable in such experiments?

As far as my attempts extend, I tried determining what conditions we should use dry mass under, but I could not find any source that talks about detailed use of dry mass (when to use it and when to not use it).

Perhaps a different way to phrase this question is to ask: when do we use mass rather than dry mass? And when do we use dry mass rather than mass? As far as my knowledge extends, dry mass is the most accurate determinant of growth that there is, given that the plants are genetically identical. This is because it ignores the possible fluctuations that may be present due to fluid mass within the plants.


What are the Methods of Measuring Microbial Growth?

There are different methods of counting microbial growth. These are based on different parameters of cells such as dry-weight and wet-weight measurement, absorbance, cell plate, density, turbidity, ATP measurement, viable count, ATPase activity and use of Coulter counter.

(a) Wet Weight Measurement:

Measuring cell mass is an easy step of cell growth measurement. A known volume of culture sample from the ferment or is withdrawn and centrifuged, Wet weight of pellets is measured by using pre-weighed filter paper. A pre-weighed filter paper of similar size is used to subtract the weight of wet filter paper. Thus wet-weight of cells is calculated.

(b) Dry Weight Measurement:

Dry weight measurement of cell material is similar to that of wet weight. Here dry weight of pre-weighed filter paper containing pellets of microbial cells is measured. Dry weight of filter paper is nullified by subtracting the dry weight of only filter paper of similar size.

Thus dry weight of microbial cells can be obtained. For example dry weight of about one million cells of E. coli is equal to 150 mg. Dry weight of bacterial cells is usually 10-20% of then- wet weight.

(c) Absorbance:

Absorbance is measured by using a spectrophotometer. Scattering of light increases with increase in cell number. When light is passed through bacterial cell suspension, light is scattered by the cells.

Therefore, transmission of light declines. At a particular wavelength absorbance of light is proportional to the cell concentration of bacteria present in the suspension.

Thus cell growth of any bacterial suspension at a particular wavelength at different intervals can be measured in terms of absorbance and a standard graph (between absorbance and cell concentration) can be prepared.

(d) Total Cell Count:

Cell growth is also measured by counting total cell number of the microbes present in that sample. Total cells (both live and dead) of liquid sample are counted by using a special microscope glass slide called Petroff-Hausser Counting Chamber.

In this chamber a grid is marked on the surface of the glass slide with squares of known area. The whole grid has 25 large squares, a total area of 1 mm 2 and a total volume of 0.02 mm 3 (1/50 mm).

All cells are counted in large square and total number per ml sample is measured. If 1 square contains 12 cells, the total number of cells per ml sample will be: 12 cells x 25 square x 50吆 3 = 1.5xl0 7 cells.

If there is dilute culture, direct cell counting can be done. However, the cell culture of high density can be diluted. Otherwise clumps of cells would be formed which would create problem in exact counting of bacterial cells.

(e) Viable Count:

A viable cell is defined as a cell which is able to divide and increase cell numbers. The normal way to perform a viable count is to determine the number of cells in the sample which is capable of forming colonies on a suitable medium.

Here it is assumed that each viable cell will form one colony. Therefore, viable count is often called plate count or colony count. There are two ways of forming plate count.

(i) Spread Count Method:

A volume of culture (0.1 ml) is spread over the surface of an agar plate by using a sterile glass spreader. The plate is incubated to develop colonies. Then the number of colonies is counted (Fig. 6.6A).

(ii) Pour Plate Method:

In this method a known volume (0.1-1.0 ml) of the culture is poured into the sterile Petri dishes. Then melted agar medium is poured and mixed gently. The plate is incubated. Colonies growing on the surface of agar are counted.

(f) Coulter Counter:

It is an electronic device. The microbial culture is directly used to count cells present in the suspension.


INTRODUCTION

There has been recent interest in the use of electrical techniques for quantifying root systems (Cao et al., 2010, 2011 Urban et al., 2011 Dietrich et al., 2012 Ellis et al., 2012). Many studies have reported good correlations between root mass and electrical capacitance, measured between an electrode inserted at the base of the stem and an electrode in the rooting substrate (e.g. Chloupek, 1977 Dalton, 1995 van Beem et al., 1998 Preston et al., 2004 Ozier-Lafontaine and Bajazet, 2005 McBride et al., 2008 Tsukahara et al., 2009). Linear relationships between root mass and electrical capacitance have been interpreted using an electrical model proposing that roots behave as cylindrical capacitors and their capacitances can be added together as though wired in parallel (Dalton, 1995). This model was tested in hydroponics by Dietrich et al. (2012), who found that the capacitance of barley (Hordeum vulgare) appeared to be determined, not by the mass of their root system, but by the cross-sectional area of roots at the solution surface. These authors also observed (i) that capacitance was not linearly related to the mass of roots in solution when root systems were partly submerged and (ii) that excising the root below the solution surface had a negligible effect on the capacitance measured. These observations are inconsistent with the model of Dalton (1995). A new model for plant capacitance was proposed by Dietrich et al. (2012), suggesting that plant tissue behaves as a continuous dielectric and, provided the capacitance of the tissue is much smaller than that of the rooting substrate, the capacitance measured in hydroponics is dominated by tissue between the solution surface and the electrode attached to the plant. The new model suggests that the measured capacitance will be inversely proportional to the distance between the plant electrode and the solution surface. This model remains to be tested in solid rooting substrates, where both the capacitance of the substrate and the contact between roots and solution are likely to be smaller than in hydroponics and will vary with the water content of the rooting substrate. In this study, the ability of the new model to explain capacitance measurements made on cereals growing in sand or in compost in the glasshouse, or in a sandy-loam soil in the field, was tested under various water regimes.


When is it unacceptable to measure dry mass in an experiment to measure growth of plants? - Biology

Principles of Ecology

Lab 2 Plant Competition

Introduction:

One of the ways in which organisms may interact is competition. In this lab, we will investigate the interaction between seedling plants. We use plants for practical reasons. The primary advantage is that the growth of plants is very plastic. This means that the size a plant reaches is largely determined by environmental factors within the general framework set by its genes (peas will not grow into trees, no matter how advantageous the environment). Factors that are possibly important include such things as nutrients, light, soil characteristics (other than nutrients) moisture, and the density of other plants (both plants of the same species and other species). The focus of this lab will be on neighboring plant density as a factor affecting plant growth.

We will grow plants in two ways in order to detect the effects of competition with other plants. We will grow an herb, sweet basil, by itself but at different densities (number of plants per pot) and measure the size of the plants that result. Since competition affects individuals, we will concentrate on the effect of competition on the average size of basil plants. We would have to do other experiments to understand the effect intraspecific competition has on basil populations. Second, we will look at the relative strength of interspecific versus intraspecific competition by growing two different species of plants, zinnia and marigold, in pots with different densities of plants per pot. We will also vary the proportion of each species in the pots, as some will be mostly zinnia and some mostly marigold. Once again, WE WILL BE INTERESTED IN THE INDIVIDUAL PLANT'S RESPONSE, MEASURED AS THE AVERAGE SIZE OF PLANTS IN A POT (FOR EACH SPECIES)

Intraspecfic Competition:

Other things being equal, does the presence of other plants from the same species affect the growth of individual plants in a predictable fashion?

You will have to fill this in with a reasonable guess based on what you know about plants and the procedures described below. BE SURE THAT YOU STATE A HYPOTHESIS AS PART OF YOUR LAB WRITE-UP. I will need to read it in order to be sure you understand what the lab is all about. This step should be done WITH CARE.

  1. Obtain a set of six pots. Put an inch or two of tape on each pot as a label and label each pot with the number and type of seeds to be planted in each plus something to identify them as your pots. Use pencil as the pots will be sprayed regularly with water and left in sunlight. Ink can fade and wash away but the pencil marks should persist.
  2. Fill each pot with soil until the soil comes to within 1 cm of the top. DO NOT TAMP THE SOIL DOWN, AS YOU CAN PREVENT THE SEEDS FROM SUCCESSFULLY GERMINATING. To settle the soil, you can tap the bottom of the pot on the table a couple of times.
  3. Plant either 2, 3, 5, 10, 18, or 34 basil seeds in each pot. After germination, the plant populations pots will be thinned to 1, 2, 4, 8, 16, 32, or 64 plants per pot. The extra seeds are to ensure we have enough plants in each pot. Do your best to space them evenly over the top of the soil.
  4. Top the seeds with more soil. Completely fill each pot with soil by gently dropping soil from your hand. (use a straightedge or piece of paper to level the top of the soil with the top of the pot). Once again, no tamping (tapping is OK).
  5. Return the pots to the flats. The plants will be watered and tended for the next several weeks.

Interspecific Competition:

Other things being equal, does the presence of other plants from other species affect the growth of individual plants differently than the presence of plants from the same species and does this difference occur in a predictable fashion?

You will have to fill this in with a reasonable guess based on what you know about plants and the procedures described below. BE SURE THAT YOU STATE A HYPOTHESIS AS PART OF YOUR LAB WRITE-UP. I will need to read it in order to be sure you understand what the lab is all about. This step should be done WITH CARE.

  1. Obtain a set of 4 pots. Put an inch or two of tape on each pot as a label and label each pot with the number and type of seeds to be planted in each plus something to identify them as your pots.
  2. Fill each pot with soil until the soil comes to within 1 cm of the top. DO NOT TAMP THE SOIL DOWN, AS YOU CAN PREVENT THE SEEDS FROM SUCCESSFULLY GERMINATING. To settle the soil, you can tap the bottom of the pot on the table a couple of times.
  3. We will use marigold and zinnia for this portion of the lab. Plant the seeds in the densities of:
    • 4 M : 4 Z
    • 4 M : 32 Z
    • 32 M : 4 Z
    • 32 M : 32 Z.

Do your best to space them evenly on the soil. Notice that each species experiences high and low amounts of interspecific and interspecific competition

Gathering the Data:

In gathering the data from the intraspecific competition experiment, you gather detailed information from each pot (number of plants, weights of plants, number of leaves, length of stem). In the interspecific experiment, all you do is count the number of plants of each species and then take the total weight of all plants in each pot.

You must weigh the plants quickly, as they will lose weight, due to water loss, as soon as you remove them from the soil. They can lose so much water that you can completely change the outcome of the work. So, work as a team. To gather the interspecific data, each team member should clip the plants from a pot and immediately weigh the plants. In the interspecific experiment, one person should clip the plants from a pot as another tallies the species each belongs to and separates the plants into piles of different species and then immediately weighs the plants from that pot as soon as they are done clipping and tallying (the tendency in previous labs was to clip all pots, separate the plants and then weigh them all at once, but this takes too much time.

  1. Count and record the number of plants in a pot and the number of leaves on each plant in the pot. Record the number of plants that produced a flower bud (if any have done so in the time we have for this exercise).
  2. Cut off all shoots at ground level.
  3. Weigh all of the plants from a pot together. Calculate the average weight by dividing this total by the actual number of plants in the pot (not the number of seeds you planted).
  4. Remove the buds and leaves from each plant stem (by pot once again!), combine them on one weighing pan, and weight them together to obtain the total leaf, bud, and stem weight in each pot. Don't worry if there are no buds. Notice that you don't have to weigh the stems, as you can get their weight by subtracting the leaf and bud total (or just the leaf total if there are no buds) from the total plant weight for the pot.
  5. Measure the length of each stem.
  6. Return the pots to the flats.
  7. Designate a member of the group who will enter the data onto a spreadsheet and email the results to the instructor by noon of the following day.
  1. Cut off all shoots from a single pot at ground level and hand them to another student who will divide them into separate species. Record the number of each that germinated.
  2. We will again ignore the below-ground portion of the plant, as we have no reliable way to separate the roots by species and it is impossible to separate complete plants without tearing apart the root ball.
  3. Weigh all of the plants from each species and divide by the total number of plants of that species in that pot to get a mean plant weight for each species in each pot.
  4. Designate a member of the group who will enter the data onto a spreadsheet and email the results to the instructor by noon of the following day.

Data Analysis: You must do the sections beginning with "To get an A . " or your maximum score will be B+

Intraspecific Competition: Please answer the following questions with graphs and tables constructed from both your data and the data from the rest of the class.

  1. You can look at the effects of intraspecific competition in several ways. The graphs, charts and calculations suggested below are intended to do just that. first you should plot the mean plant size versus density of plants in the pot and do a second plot of total plant biomass in the pot verss plant density in the pot:
    • How does competition affect the mean weight of plants in a pot?
    • What is the relationship between total biomass and density? Compare this to the mean weights. Does total weight tell you anything about intraspecific competition?
  2. Plot the average stem length versus number of plants in the pot(include standard deviation error bars).
    • What does this plot tell you about intraspecific compeition?
  3. Do plants change the proportion of biomass allocated to leaf, shoot and root as density increases? Since the number of plants in a pot was not strictly controlled, we can not make a sensible bar chart. Use a scatter plot of density versus the three proportions (stem, leaf, and bud) and examine the graph to draw your conclusions.
    • How does density affect the proportional allocation of biomass to the various parts of the above-ground portion of the plant?
  • To get and A, you must do the following analysis. A long time ago, Kira et al. (1953) proposed that the relationship between mean plant weight and density could be described as:

where w = mean plant weight and p = density. K and a are constants used to fit this relationship to different plants and must be estimated from the data. We can do this, but must first linearize the relationship by taking the log of both sides:

This equation is in the form of y = mx + b, the standard linear equation (the slope is negative here), If you plot this, with your y values = log(w) and your x values = log(p), the points should form a line. You will, of course, not get a perfect line, but you can estimate the real relationship by estimating the line. Your spreadsheet program will do this estimation for you if you ask for a linear trend line (be sure to get the equation for the line). Use the equation to get your estimates of a and K. Use a and K to get estimated mean plant sizes for your densities. Add the expected data to the graph of mean plant wt. versus density as a second variable (you may want to add more densities to get a smooth line). How does the expected line agree with the observed data?

There are many possible ways to analyze the data from this portion of the lab. You want to see how the average size of a plant changes as its neighbors are more and more from another species. One of the most visually interpretable methods was devised by De Wit (De Wit, 1961 Harper, 1967). The diagrams are called De Wit replacement plots and are easy to make. Make a separate graph for each species. Plot the average plant weight for each density (total density here is approximately 10, 55, and 100 plants). There should be two points at the intermediate density, one for the 5:50 pot and one for the 50:5 pot. Notice that this assumes that all of the seeds have germinated, an unlikely occurrence. Your plot will likely have different densities and, on the X axis, you should plot the total number of plants you actually measured, not the number of seeds planted. In the example below, dotted lines indicate the interspecific effect and solid lines the intraspecific effect (THIS IS NOT SO EASY TO SEE SO BE SURE THAT YOU UNDERSTAND WHY THIS IS SO). Look hard at the graph below and read the interpretation of in underneath.

The solid line from the dot at 20 plants per pot to the dot labeled 100M:10Z represents the change in marigold size when the number of marigold seeds go from 10 to 100 while the number of clover seeds remains constant at 10. So, the solid line is one estimate of the effect of interspecific competition on marigold. The dotted line above it is just the opposite. Here, it is the marigold plants that are constant at 10 in each and the zinnia goes from 10 to 100 plants in the pot. So the dotted line represents the interspecific effect of zinnia on marigold seedling weight. In this diagram, the average weight of Marigold plants is reduced more by being in a pot with 110 plants if 100 of them are marigolds than if 100 of them are zinnias. I can see this because the slope of the intraspecific (solid) line is more negative than the slope of interspecific (dotted) line. Remember what slope means. The second set of lines (from 100:10 to 200) will give you the same conclusion. Plot your data and draw conclusions from what happened in your pots (this is hypothetical here).

What you must do, from the two graphs from your data, is to draw a conclusion about which has a greater effect interspecific competition or intraspecific competition? Of course, your conclusion must be reasonably argued FROM THE DATA .

Thinking about the experiment:

  1. What is the most likely mechanism by which one plant affects its neighbors in the intraspecific experiment? Mechanism here refers to the actual way in which plants affect one another's growth. There are many possibilities and explain why you made your choice.
  2. Is the mechanism of interaction likely to be the same in both the intra- and interspecific experiments?
  3. Design an experiment to test your idea from Question #1. Describe it in enough detail to show that you have drawn on what you have learned in other classes in order to develop this experiment.
  4. Is the mechanism you have chosen an instance of exploitation competition or one of interference competition? Whichever you choose, describe a possible mechanism of interaction that belongs to the opposite type of competition.

Classic References:

Black, J. N. 1960. An assessment of the role of planting density in competition between the red clover (Trifolium pradense L.) and lucerne (Medicago sativa L.) in the early vegetative stage. Oikos 11:26-42.

Clements, F. E., J. E. Weaver, and H. C. Hansen. 1929. Plant Competition. Carnigie Institute, Washington, D. C.

de Wit, C. T. 1961. Space relationship within populations of one or more species. Society of Experimental Biology Symposium 15:314-329.

Harper, J. L. 1967. A Darwinian approach to plant ecology. Journal of Ecology 55:247-270.

Kira, T., H. Ogawa, and N. Sakazaki. 1953. Intraspecific competition among higher plants. I. Competition-yield-density interrelationship in regularly dispersed populations. Journal of the Polytechnic Institute of Osaka City University D 4:1-16.

Link to dataset if a greenhouse disaster occurs:

Materials (startup week in bold):

Sweet Basil, Zinnia and Marigold Seeds, Pots (11 x #of groups), Straight edges, Soil, Paper Tape, Scissors or razors, Balances, Rulers, Buckets, Soil Storage Container


Determination of Moisture Content in Soil

Soils normally contain a finite amount of water, which can be expressed as the “soil moisture content.” This moisture exists within the pore spaces in between soil aggregates (inter-aggregate pore space) and within soil aggregates (intra-aggregate pore space) (Figure 1). Normally this pore space is occupied by air and/or water. If all the pores are occupied by air, the soil is completely dry. If all the pores are filled with water, the soil is said to be saturated.


Figure 1. Pore space in soil.

Principles

In outdoor natural environments, water is added to soil via rainfall or deliberate irrigation of plants. In either case, soil moisture increases as more pores become filled with water at the expense of air. If all the pores become filled with water, excess water will now leach downward (Figure 2) through continuous soil pores, until the rain or irrigation ceases. Leaching will continue until the water films within the pores are held by the surface tension of soil colloids against the force of gravity. Such a situation is referred to as the soil being at “field capacity” with respect to soil moisture. A soil at field capacity has pores partially filled with air, surrounded by soil moisture films. Normally a soil at field capacity is optimal for plant growth and aerobic soil microorganisms, since both air and water are available. In contrast, a saturated soil will create waterlogged anaerobic conditions that can kill plants and suppress aerobic soil microbes, while stimulating anaerobic microbes.


Figure 2. Nutrients leaching in soil.

Consider a sample of moist soil within a container such as a beaker. The weight of the moist soil consists of the weight of the dry soil particles plus the weight of the water within the soil. If more water is added to the soil, the wet weight of the soil increases. The dry weight of the soil particles within the sample is fixed i.e., one weight which is the dry weight. In contrast, there are an infinite number of wet weights, depending upon how much water is added to the soil. Because of this, when doing lab experiments with soil, the moisture content of the soil is normally expressed on a dry weight basis, because the dry weight is constant over time, whereas the moist or wet weight can change over time. When expressing the results of an experiment such as the nutrient content of a soil, use of the dry weight basis provides standardization of the final result.

Procedure

  1. Weigh both of the aluminum dishes.
  2. Aliquot approximately 50 g of moist soil into each aluminum dish and reweigh the dishes. Hence, the moist weight of the soil sample is now known.
  3. Dry the soil overnight at 105 °C in the oven.
  4. Remove the dishes from the oven and allow them to cool.
  5. Reweigh the dishes plus the oven dry soil. Now the weight of the dry soil is known.

The amount of water held in soil is an important component of biological and ecological processes, and is used in applications such as farming, erosion prevention, flood control, and drought prediction.

Soils typically contain a finite amount of water, which can be expressed as the soil moisture content. Moisture exists in soil within the pore spaces between soil aggregates, called inter-aggregate pore space, and within pores in the soil aggregates themselves, called intra-aggregate pore space. If the pore space is occupied entirely by air, the soil is completely dry. If all of the pores are filled with water, the soil is saturated.

The measurement of the amount of water held within the soil, or the soil moisture content, is essential to the understanding of soil characteristics and the types of plants and microorganisms that reside in it.

This video will introduce the basics of soil moisture content, and demonstrate the procedure for determining moisture content in the laboratory.

In outdoor environments, water is added to soil naturally through rainfall or deliberately with the irrigation of plants. As the pores in the soil become filled with water at the expense of air, the soil moisture increases. When all of the pores are filled with water, the soil is saturated. If the soil at the surface is saturated, excess water will leach downward through pores into deeper soil. Leaching continues until there is not enough water to saturate all of the pore space. At this point pores contain some air and thin films of moisture. The water films within the pores are held by the surface tension of soil colloids, thus water stops leaching.

After leaching stops, and excess water has drained from the soil, the soil is described as being at field capacity. Soil at field capacity has pores that are partially filled with air, surrounded by films of moisture. Soil at field capacity is optimal for plant growth and aerobic soil microorganisms, since both air and water are available. In contrast, saturated soil, where all pores are filled with water, will create an anaerobic environment that can kill plants and suppress aerobic soil microbes.

The mass of moist soil consists of the mass of the dry soil particles, plus the mass of the water within the soil. The dry mass of the soil particles is fixed, whereas the amount of water within moist soil can vary. Therefore, moisture content is calculated on a dry basis, rather than a total mass basis, to ensure consistency. The moisture content of soil is described as the ratio of the mass of water held in the soil to the dry soil. The mass of water is determined by the difference before and after drying the soil.

The following experiment will demonstrate how to measure soil moisture content in the laboratory using these principles.

To begin, collect soil samples and transfer them into the laboratory. Samples of soil can be collected in the field using a soil auger, or a trowel. Use of a soil auger allows for the soil to be sampled to specific depths. Transfer them into the laboratory. Weigh two aluminum dishes, and accurately record the weight of each dish. Aliquot approximately 20 g of the moist soil into each aluminum dish, then reweigh the dish. Subtract the weight of the empty dish from the full dish to acquire the moist soil weight.

Next, dry the soil overnight in an oven set to 105 °C. On the next day, carefully remove the soil samples from the oven using tongs. Place the soil samples on the bench top to cool. When the dry soil samples are cool, reweigh them and record the total weight. Subtract the weight of the aluminum dish, and record the dry soil weight.

Calculate the moisture content of the soil by subtracting the weight of the dry soil from the weight of the moist soil, and then dividing by the weight of the dry soil.

Although the measurement is simple, it is important to determine soil moisture content in order to better understand soil characteristics.

Soil moisture content plays a large roll in environmental concerns, especially when considering soil runoff that may contain fertilizers and pesticides. In this example, soil runoff was analyzed using a simulated rainfall study in order to determine the retention of a compound in moist soil.

Soil, containing urea, was packed into soil boxes and assembled under a rainfall simulator. Soil runoff was collected, and the concentration of urea in the runoff water calculated. The amount of urea in the soil runoff was higher for soils that had higher moisture content, indicating that urea is better absorbed in drier soil, than in moist.

The fate of chemicals in soil can also be analyzed by direct pore water sampling, using a lysimeter, as shown in this example. In this experiment, lysimeters, or long metal tubing, were installed in soil with turf grass to analyze pore water in vegetative soil.

The pore water sampler was then installed, and water pumped from the lysimeter after applying chemicals to the soil. The collected water was then analyzed, and the concentration of applied chemicals correlated to soil depth and moisture content.

The results demonstrated that the concentration of the herbicide monosodium methyl arsenate, or MSMA, was the highest in the top 2 cm of soil.

You've just watched JoVE's introduction to soil moisture content. You should now understand how to accurately measure soil moisture content in the laboratory. Thanks for watching!

Results

Calculate the soil moisture content for each of the replicate samples using the following equation:

% moisture content (MC) =

∴ % MC =

With the addition of 5 g of water, new M = 107 and D still equals 90

∴ % MC =

Applications and Summary

Knowledge of the moisture content of a soil on a dry weight basis is useful in a number of ways. For example, if the experiments are conducted with soil that should be amended with a known concentration of ammonium fertilizer (for example 50 μg/g), then the moisture content on a dry weight basis must be determined. If the calculation was completed on a wet weight basis, the amount of fertilizer to be added would depend on the moisture content (and therefore the moist weight) of the soil sample. Likewise, if potted plants are considered, the moisture content must be known in order to make sure that the soil isn’t too dry (not enough moisture for plant growth) or too wet (waterlogged and anaerobic). In a field situation, knowledge of the soil moisture content can prevent excess irrigation and leaching of soil nutrients.

Transcript

The amount of water held in soil is an important component of biological and ecological processes, and is used in applications such as farming, erosion prevention, flood control, and drought prediction.

Soils typically contain a finite amount of water, which can be expressed as the soil moisture content. Moisture exists in soil within the pore spaces between soil aggregates, called inter-aggregate pore space, and within pores in the soil aggregates themselves, called intra-aggregate pore space. If the pore space is occupied entirely by air, the soil is completely dry. If all of the pores are filled with water, the soil is saturated.

The measurement of the amount of water held within the soil, or the soil moisture content, is essential to the understanding of soil characteristics and the types of plants and microorganisms that reside in it.

This video will introduce the basics of soil moisture content, and demonstrate the procedure for determining moisture content in the laboratory.

In outdoor environments, water is added to soil naturally through rainfall or deliberately with the irrigation of plants. As the pores in the soil become filled with water at the expense of air, the soil moisture increases. When all of the pores are filled with water, the soil is saturated. If the soil at the surface is saturated, excess water will leach downward through pores into deeper soil. Leaching continues until there is not enough water to saturate all of the pore space. At this point pores contain some air and thin films of moisture. The water films within the pores are held by the surface tension of soil colloids, thus water stops leaching.

After leaching stops, and excess water has drained from the soil, the soil is described as being at field capacity. Soil at field capacity has pores that are partially filled with air, surrounded by films of moisture. Soil at field capacity is optimal for plant growth and aerobic soil microorganisms, since both air and water are available. In contrast, saturated soil, where all pores are filled with water, will create an anaerobic environment that can kill plants and suppress aerobic soil microbes.

The mass of moist soil consists of the mass of the dry soil particles, plus the mass of the water within the soil. The dry mass of the soil particles is fixed, whereas the amount of water within moist soil can vary. Therefore, moisture content is calculated on a dry basis, rather than a total mass basis, to ensure consistency. The moisture content of soil is described as the ratio of the mass of water held in the soil to the dry soil. The mass of water is determined by the difference before and after drying the soil.

The following experiment will demonstrate how to measure soil moisture content in the laboratory using these principles.

To begin, collect soil samples and transfer them into the laboratory. Samples of soil can be collected in the field using a soil auger, or a trowel. Use of a soil auger allows for the soil to be sampled to specific depths. Transfer them into the laboratory. Weigh two aluminum dishes, and accurately record the weight of each dish. Aliquot approximately 20 g of the moist soil into each aluminum dish, then reweigh the dish. Subtract the weight of the empty dish from the full dish to acquire the moist soil weight.

Next, dry the soil overnight in an oven set to 105 °C. On the next day, carefully remove the soil samples from the oven using tongs. Place the soil samples on the bench top to cool. When the dry soil samples are cool, reweigh them and record the total weight. Subtract the weight of the aluminum dish, and record the dry soil weight.

Calculate the moisture content of the soil by subtracting the weight of the dry soil from the weight of the moist soil, and then dividing by the weight of the dry soil.

Although the measurement is simple, it is important to determine soil moisture content in order to better understand soil characteristics.

Soil moisture content plays a large roll in environmental concerns, especially when considering soil runoff that may contain fertilizers and pesticides. In this example, soil runoff was analyzed using a simulated rainfall study in order to determine the retention of a compound in moist soil.

Soil, containing urea, was packed into soil boxes and assembled under a rainfall simulator. Soil runoff was collected, and the concentration of urea in the runoff water calculated. The amount of urea in the soil runoff was higher for soils that had higher moisture content, indicating that urea is better absorbed in drier soil, than in moist.

The fate of chemicals in soil can also be analyzed by direct pore water sampling, using a lysimeter, as shown in this example. In this experiment, lysimeters, or long metal tubing, were installed in soil with turf grass to analyze pore water in vegetative soil.

The pore water sampler was then installed, and water pumped from the lysimeter after applying chemicals to the soil. The collected water was then analyzed, and the concentration of applied chemicals correlated to soil depth and moisture content.

The results demonstrated that the concentration of the herbicide monosodium methyl arsenate, or MSMA, was the highest in the top 2 cm of soil.

You've just watched JoVE's introduction to soil moisture content. You should now understand how to accurately measure soil moisture content in the laboratory. Thanks for watching!


Determination of Specific Leaf Area and Leaf Area-leaf Mass Relationship in Oil Palm Plantation

Specific leaf area (SLA), the ratio of leaf area to leaf mass is the most important determinant of oil palm growth, which is used in growth monitoring of oil palm and many crop simulation model s to estimate total leaf area. Leaf dry weight and leaf area were determined by destructive methods in oil palm plantation. The objective of this study was to obtain suitable linear model for estimation of leaf area and calculation SLA of oil palm plantation with less error of estimation. The SLA was plotted on frond number and found that SLA was decrease systematically with time as the frond mature. In this study we found that the Leaf dry weight was strongly correlated (R 2 =0.96-0.98) with leaf area in both linear and non-linear regression. The leaf mass were regressed on leaf area using both linear and non-linear model and found following relationship: Leaf Mass=Leaf Area/99 (R 2 =0.96) Leaf mass were calculated from rectangular leaf area by above linear equation. Leaf Mass=0.0087 (Leaf Area) 1.027 (R 2 =0.98) Leaf mass were calculated from rectangular leaf area by above non-linear equation. Actual leaf area (individual or whole of the oil palm plantation) were calculated from leaf dry weight by following linear and non-linear regression equation: Actual Leaf Area=78.89 x Leaf Mass (R 2 =0.97) Actual Leaf Area=80.926 (Leaf Mass) 0.977 (R 2 =0.98) Its also found that the calculated leaf area and measured leaf area was strong linear relationship (R 2 =0.98).

M.A. Awal, Wan Ishak , J. Endan and M. Haniff , 2004. Determination of Specific Leaf Area and Leaf Area-leaf Mass Relationship in Oil Palm Plantation. Asian Journal of Plant Sciences, 3: 264-268.

Leaf area and leaf mass relationships can be expressed by the Specific leaf area (SLA) (cm 2 g -1 or m 2 Kg -1 ), which is the ratio leaf area to leaf dry weight. According to Barden [1] SLA has been related to leaf structure, growth and net photosynthesis. Also SLA is used in crop simulation model s to estimate total leaf area or dry weight [2] . Leaf area and specific leaf area are important parameters in many agronomic and ecological processes, including photosynthesis, transpiration and field energy balance, but can be difficult and expensive to measure [3] . It is used to estimate total leaf area at various stages of growth and many crop model to predict leaf area from leaf dry weight or vice versa. SLA also can be used in conjunction with leaf area to estimate leaf mass for nutrient balance calculations and growth estimates. In most crops leaf area is defined by the leaf area index LAI. This term expresses the area of the aboveground plant components such as leaves, branches and fruit per unit area of ground in m 2 m -2 [4] .

In oil palm, it is difficult and time consumes to measure total leaf area in various stages. But it can be easily estimated by SLA. From SLA model, any oil palm plantation manager could be estimate total leaf area of the plantation. Many crop models calculate either leaf area or leaf dry weight and use SLA to determine the value of the other variable. Some scientists have assumed a constant SLA for leaves after full expansion [5,6] .

The intrinsic photosynthetic capacity of palms depend on the leaf structural characteristics, such as leaf thickness, size and arrangement of mesophyll cells that determine the amount of photosynthetic tissue per unit leaf area. Also SLA indicates leaf thickness of the leaflet. Therefore, parameters such as specific leaf area (SLA), specific leaf dry weight (SLDW, dry weight per unit leaf area) or the total content of chlorophyll per unit area of leaf (SPAD) are considered good indicators of the strength of the photosynthetic tissue [7] . However, These parameters are important for forest and agricultural research as well for crop management practices. The inverse of SLA, specific leaf mass or specific leaf dry weight, has been positively correlated with leaf water use efficiency among alfalfa ( Medicago sativa L.) cultivars by Gutschick [8] who reasoned that leaves with high specific leaf mass are cooler under a given radiation load due to higher stomatal conductance and lower water vapor pressure deficit. Leaf area and leaf mass are closely related to light interception, photosynthesis, transpiration, growth rate and furthermore to yield [9] and Charles-Edwards [10] has also shown a positive correlation between SLA and light use efficiency for several species. Thus SLA is an important crop parameter to estimate.

The objectives of this study were: to determine the SLA from destructive sampling and direct measurements of leaf area and mass in oil palm and determine total leaf mass of oil palm from leaf area measurements. Also to test a statistical model that calculates SLA of individual leaves from their dry mass.

Experiments were conducted in experimental plot at MPOB during 2003 to estimate the leaf area, leaf mass and specific leaf area. The first experiment was conducted during May 2003. The second experiment was performed at same plot during October 2003.

Study site: Measurements were made in Malaysian Palm Oil Board (MPOB) ENOVECY research plot. MPOB is situated about 30 km north from Kuala-Lumpur, Latitude 2° 58 / 0.36 // N, Longitude 101° 44 / 26 // E) at an average altitude of 66.5 m from sea level. Agronomy division at MPOB in 1998 planted the plantations. We Considered 5-6 years old uniform palm Tenera (D x P) for this study.

For leaf area measurement, two methods were used in this study

Manually measurement
Measurement by Portable Leaf Area Meter (Li-3000A Leaf Area Meter)

LI-3000A leaf area meter: The LI-3000A combines an easy to use, microprocessor controlled readout console with the proven scanning technology of the LI-COR LI-3000 sensor head to provide a powerful system for portable non-destructive leaf area measurements. The LI-3000A utilizes an electronic method of rectangular approximation to provide one mm 2 resolution. The readout console logs leaf area, leaf length, average width and maximum width as the scanning head is drawn over a leaf. Files can be viewed on the display or output through the RS-232C interface to a computer or printer. For large numbers of detached leaves, the LI-3000A can be used with the LI-3050A Transparent Belt Conveyor Accessory for greater measurement efficiency.

Measurement of leaf area: For measurement of leaf area, two types of procedure were chosen. The first one was for selected palm and other was for randomly selected palms.

Measurement of leaf area by manually in selected palm: Five palms were chosen randomly in the plantation site and frond 1, 9, 17 and 25 were chosen from each palm. The chosen leafs (frond 1, 9, 17 and 25) ware cut at the petiole level. After cutting the frond, that was brought in cool room as soon as possible for prevent of shrinkage. The length of the rachis was measured and cross-section of the petiole at appropriate point was also measured at same time. Count the leaflet in both side of rachis. Rachis length was divided in to equal ten sections. Leaflets were chosen both side of the rachis. An upper and lower leaflet with good edges was taken from the middle of each section. For both side leaflets numbered 1 to 10 on the underside of the leaflet by using permanent marker. Total twenty leaflets (each side ten leaflets) were placed according to numbering on the table. For measurement, a steel measuring scale was taken and carefully measured length (L) and middle width (W) in cm of each leaflet.

The area of each leaflet was measured by following equation:

Here, Ar represents the rectangular leaf area.

This rectangular leaf area used many experiment as well as growth monitoring of the oil palm. Where as, actual leaf area needs for many ecological modeling and LAI. So for actual leaf area calculation we used following relationships

But in this case, for leaf area measurement, leaf length and middle width were measured. That means the yield leaf area was rectangular leaf area and that was slightly higher than actual leaf area.

Measurement of leaf area by manually in random palms: In this case, several palms were chosen for investigate. Under this investigate, different size of leaflet were chosen from different fronds. Total 50 leaflets were cut for this purposes including small leaflet and large leaflet from whole of the plantation. After cutting the leaflet, it was put in plastic bag and brought in cool room as soon as possible for prevent of shrinkage. Measurement was performed by stainless steel scale according to above procedure.

Measurement of leaf area by leaf area meter: After chosen the leaflets, the leaflets were dissected from petiole, numbering and placing of the leaflet on the table as same way as manual measurement. After placing on the table, each leaflet area was measured by portable Leaf Area Meter. For more information, we record leaf area, maximum width, average width and maximum length. Recorded maximum length was compared by manually measured length and then adjusts the length of the Leaf Area Meter. Finally, more perfect results were records in the data sheet. Before measurement, Leaf Area Meter was calibrated to correct the leaf area of each sample.

Determination of leaf dry weight: All marking leaflet of experiment one and two were put carefully in the special paper bag and again mark on the bags. All bags with leaflets were oven dried at 70°C for 72h to obtain the dry mass. After drying, a precision weighing balance was used to weight of the leaflets and records the results on data sheet. A statistical analysis was done to find out a relationship between leaf areas and leaf mass by quadrant and for SLA model.

Relationship between specific leaf area and frond ages: A good linear relationship was found between the frond ages and specific leaf area. Figure 1 shows that SLA was decreased with the frond ages.

The SLA was comparatively higher at younger frond respect to mature frond. Maximum SLA 88.19 cm 2 g -1 found in frond 1, palm 1 and minimum SLA 76.39 cm 2 g -1 in frond 25, palm 3. SLA decreases systematically with time as the leaves mature, but increase systematically with depth in the canopy as the light available for leaf development and light interception decreases. There is also evidence that for a given light environment, species with leaves of higher SLA will have a higher relative growth rate.

Relation between leaf mass, leaf area and SLA: Figure 2 shows that leaf mass and leaf area relation was linear. In this study leaf area was measured directly by apply model (1). Maximum leaf area and corresponding leaf mass was 379.96 cm 2 and 3.76 g. Whereas minimum leaf area and corresponding leaf mass 24.88 cm 2 and 0.26 g. Maximum and minimum SLA was 123.54 cm 2 g -1 and 81.69 cm 2 g -1 . From this observation we proposed a mathematical linear model for calculating leaf mass from leaf area as follows

Lm is the mass of leaflet and
La is the rectangular leaf area of the leaflet

Data indicated a high degree of association (R 2 =0.96) and the low standard error of estimate of coefficient (0.00028) suggested that the relation accurately estimates oil palm leaf mass, which was used for SLA estimation. From non-linear relation, we found a non-linear model for calculating leaf mass from leaf area.

Data indicate a high degree of association (R 2 =0.98) and the low standard error of estimate of coefficient (0.0009). In this case, determination leaf mass depends on accurate measurement of leaf length and leaf middle width.

From Fig. 3 shows that leaf mass and leaf area relation was linear. From this experiment, maximum leaf area and mass was 296.26 cm 2 and 3.76 g. Minimum leaf area and mass was 20.6 cm 2 and 0.26 g. From this observation we proposed a mathematical linear model for calculating actual leaf area from leaf mass as follows:

L m is the mass of leaflet and
L ac is the actual leaf area of the leaflet

Data indicate a high degree of association (R 2 =0.97) and the low standard error of estimation were 0.169. The standard error of estimate of coefficient was (0.00029).

In this case, determination actual leaf area depends on accurate measurement of leaf dry weight. From non-linear relation, we found a non-linear model for calculating actual leaf area from leaf dry weight.

Data indicate a high degree of association (R 2 =0.98) and the low standard error of estimate of coefficient (0.0009). It is important noting that in oil palm, the relation between leaf area and leaf mass hardly changes and SLA also remain constant with growth of tree. This does not agree with Payne et al . [3] finding that the SLA gradually increase with decreased of leaf mass. However the data present here support the conclusions of Aase [11] for the fifth stage of growth of winter heat and results of Ramose et al . [12] for all cultivars of winter barley.

The plot of SLA vs. leaf area and leaf mass (Fig. 4) was more scattered and non-linear relationship. Figure 4 shows that the SLA varied with leaf area or leaf mass. Large leaf area represents more SLA and also more leaf dry weight represents more SLA. Because of that from Fig. 2 and 3 we found that leaf area and Leaf mass relationship was linear. It means large leaf area represents more leaf mass and specific leaf area (SLA) remains constant during the plants development and there is no significant variation with time. Because of that the leaf area increased means increased its length, width and also thickness. So that increased its mass. According to Blackman [13] the environmental factor, such as light and temperature were modifying the SLA. Acock [14] . Found that SLA differed accordingly to the position of the leaf area on the plant. The distal or youngest leaf sampled had the highest SLA, suggesting that leaf area of a developing leaf stabilizes before its dry weight. This study results strongly support these experimental results.

From the present study it can be concluded that leaf dry weight for oil palm plantation gives good estimate of the actual leaf area, where light and temperature were not much changes. As the same way, Leaf mass can be calculated from simply measured rectangular leaf area.

SLA appears to be constant with increasing leaf area or leaf dry weight. SLA varied with environmental factors like light and temperature. From Fig. 1 it was clear that SLA varied with frond maturity as well as frond position. Younger frond (frond-1) situated at the upper portion of the tree and hence received more light. But in lower frond (frond-25) received comparatively low light intensity. Because light intensity decreased rapidly with increased the depth of canopy. Probably low light intensity was responsible for less SLA in lower frond. For measurement of SLA in oil palm plantation, it was important to accurate calculation of leaf area and leaf mass rather then used crop model.

The authors thank Mr. Mohd. Hudzari b. Razali for harvested frond from oil palm trees removal of leaf from frond and collected data. We appreciated the cooperation of Biological Division, MPOB for helping us by logistic support as well as providing to Laboratory facilities and permission to harvest the fronds.

2: Reddy, V.R., B. Acock, D.N. Baker and M. Acock, 1989. Seasonal leaf area-leaf weight relationships in the cotton canopy. Agron. J., 81: 1-4.

3: Payne, W.A., C.W. Wendt, L.R. Hossner and C.E. Gates, 1991. Estimating pearl millet leaf area and specific leaf area. Agron. J., 83: 937-941.
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4: Wells, J.M., 1990. Some indirect methods of estimating canopy structure. Remote Sens. Rev., 5: 31-43.

5: Stapleton, H.N., D.R. Buxton, F.L. Watson, D.J. Nolting and D.N. Barker, 1973. Cotton: A computer simulation of cotton growth. Arizona Agric. Exp. Stn. Tech. Bull., 124: 44-44.
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6: Gutschick, V.P., 1988. Optimization of specific leaf mass, internal CO2 concentration and cholophyll content in crop canopies. Plant Physiol. Biochem., 26: 525-537.

7: Ma, L., F.P. Gardner and A. Selamat, 1992. Estimation of leaf area from leaf and total mass measurments in peanut. Crop Sci., 32: 467-471.
CrossRef |

8: Charles-Edwards, D.A., 1982. Physiological Determinants of Crop Growth. Academic Press, Sydney, New York.

9: Aase, J.K., 1978. Relation between leaf area and dry matter in winter wheat. Agron. J., 70: 563-567.

10: Ramos, J.M., L.F.G. del Moral and L. Recalde, 1982. Dry matter and leaf area relationships in winter barley. Agron. J., 75: 308-310.
CrossRef |

11: Acock, M.C., C.S.T. Daughtry, G. Beinhart, E. Hirschmann and B. Acock, 1994. Estimating leaf mass from light interception measurements on isolated plants of erythroxylum species. Agron. J., 86: 570-574.
Direct Link |

12: Brown, L.G., J.W. Jones, J.D. Hesketh, J.D. Hartsog, F.D. Whisler and F.A. Harries, 1985. Computer simulation of cotton growth and yield. COTCROP, Mississippi Agriculture Forest Experiment Station Information Bulletin, pp: 69.

13: Haniff, M., 1999. Annual progress report. Biology division, MPOB, Bangi, Malaysia.


Seed Germinator

What goes on underground when seeds are sprouting? Make yourself a window into the process of plant development.

Tools and Materials

  • Quick-germinating seeds, such as radish or Wisconsin Fast Plants®
  • Water
  • Paper towel or coffee filter
  • Scissors
  • Pencil
  • Large petri dish with lid, or an old CD case with clear sides (if you’re using a CD case, open it and remove the plastic insert that holds the CD, being careful not to break the case)
  • Lidless, straight-sided plastic container wide enough to set the petri dish or CD case inside, on its edge, as shown in the photo
  • Two rubber bands big enough to fit around the open container
  • Metric ruler with millimeter markings
  • Magnifying glass

Assembly

  1. Soak the seeds overnight in water.
  2. Set aside the top of the petri dish, or open the CD case. Cut the paper towel (or coffee filter) to fit inside.
  3. With a ruler and pencil, draw a straight line across the middle of the paper towel. Lay the marked-up paper in the bottom of the dish (or inside the CD case) so the line sits horizontally across the center. If you’re using a CD case, be sure the hinged edge is at the top or side (not bottom).
  4. Pour a little water into the dish to wet the paper towel. Smooth out any bubbles and tip out any extra water not absorbed by the paper. Later, when you stand the dish on its edge, the wet paper should remain stuck to the inside of the dish or CD case.
  5. Place 6 to 10 seeds on the paper towel, evenly spaced along the reference line. Then put the lid on the petri dish, or close the CD case.
  6. Stretch the rubber bands, set close to one another, around the center of the straight-sided plastic container (see photo below). Stand the petri dish (or CD case) between the rubber bands, and adjust the setup so it’s secure, standing on edge, upright in the container. With gentle handling, the seeds should stick to the moistened paper towel. If they move, put them back in their places on the line.

To Do and Notice

Check on your seeds once or twice a day, and notice what changes or emerges (see photo below). (It’s fine to open the seed germinator just handle it carefully so the seeds don’t move.) Do shoots with green tips emerge first, or do white roots emerge first? Do each seed’s roots and shoots sprout in the same direction, or in different directions? Use a magnifying glass to examine the growing structures in more detail. How do they change over time?

Measure the growth of the roots and shoots over time. You may want to collect data to graph average root length vs. time, and average shoot length vs. time. (Note that it’s helpful to measure time in total elapsed hours, rather than days.) Which grows faster, the shoots or the roots?

What's Going On?

Inside a seed is the embryo of a plant, plus a food source for that embryo, all contained within a protective seed coat. Here, you can observe seed germination, in which the embryo begins to digest the food and grow into a seedling. While this process usually happens in soil, the key component for germination is water.

At appropriate temperatures, most seeds begin their germination by absorbing water through a tiny hole in the seed coat. The moisture starts the metabolic processes of the embryo that’s contained within the seed. When hydrated by absorption of water, enzymes in the seed are activated. They begin digesting the food stored inside to generate energy for the embryo’s growth.

The developing root emerges from the seed first. As the root grows longer and thicker, it develops tiny root hairs, which help the developing plant take up water and nutrients. Shoots with pale-green leaflike structures emerge after the roots. Eventually, these leaves will turn a deeper green color and begin to photosynthesize, capturing and storing light energy and carbon dioxide from the air.

Photosynthesis in leaves supplies the plant with the energy and matter it needs to grow. Newly germinated seedlings, however, are not yet photosynthesizing. Instead, in early stages of growth, the embryo digests and assimilates the energy and matter from the food present in the seed. Depending on the type of seed, this food store contains a mixture of proteins, fats, sugars, and starches. This stored food isn’t just important to the developing plant embryo it’s also important to human diets. About 45 percent of the calories humans consume globally comes from seed grains like rice, wheat, and corn.

A common misconception is that plants get their mass from soil. In this soil-free experiment, you can prove to yourself that plants don’t strictly require soil to grow. In fact, many plants grow very well hydroponically in water cultures, as long as the appropriate nutrients such as nitrogen, phosphorous, and potassium are provided. Your seedlings will eventually need more space than the seed germinator can provide, but given the right lighting conditions, they’ll begin to photosynthesize, accumulating mass from the carbon dioxide in the air and the water you provide.

Going Further

This seed germinator makes it easy to design and perform experiments to determine the materials and conditions seeds need in order to germinate and grow. The effects of temperature, light levels, and water conditions (such as pH or salinity), as well as the presence or absence of various nutrients are all factors you can investigate. Experimenting with the position and lighting of the seed germinator can help you determine the conditions necessary for roots to grow down, and for shoots to grow up. Do roots sense gravity? Do they sense light? Or are they affected by other factors?

Teaching Tips

As noted in the What’s Going On? section above, this experiment can be used to begin investigating where a plant’s mass comes from. To do this, weigh the seeds before you soak them. Allow the seedlings to grow under good light conditions for several days, so they begin to accumulate mass through photosynthesis. Then take them out, let them dry, and check their mass again. Comparing the initial mass of the seeds to the dry mass of the germinated seedlings can help show students that a plant’s mass does not come from soil or water.


How to grow your own science experiment

Does fertilizer make plants bigger? My editor and I grew radishes to find out. Spoiler: Ours did not look this good.

Nastco/iStock/Getty Images Plus

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December 9, 2020 at 6:30 am

This article is one of a series of Experiments meant to teach students about how science is done, from generating a hypothesis and designing an experiment to analyzing the results with statistics. You can repeat the steps here and compare your results — or use this as inspiration to design your own experiment.

Sometimes when you tend a garden, your plants end up looking oddly sad. Maybe they’re short and stubby, or not as leafy as you’d like. The first thing some people might suggest is to add a little fertilizer to make your plants bigger and taller. But will fertilizer do that? Here’s an experiment to find out.

Explainer: The fertilizing power of N and P

Plants are marvels. Using carbon dioxide, light and water, they can make sugar out of (almost) thin air. “Most of a plant is made from carbon dioxide,” explains Jessica Savage. “A lot of times people think the plant grows or is built out of things from the soil. But it’s growing out of the air.” As a botanist, Savage studies plants. She works at the University of Minnesota in Duluth.

Plants can’t quite survive on air alone. They do need a few other elements. For example, the backbone of DNA — the molecule with the plant’s genetic instructions — has phosphorus atoms in it. So does ATP, the chemical that helps transfer energy around a cell. Proteins — molecules that do much of a cell’s work — need nitrogen atoms.

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Usually, plants get nitrogen and phosphorus from the soil. Some plants are known as nitrogen-fixing. They can pull nitrogen from the air and transform it into nitrogen-containing molecules that plants can use. But most plants can’t do this. They have to rely on other plants or fungi to transform nitrogen for them. They also have to get phosphorus in the form of phosphate (phosphorus bound to four oxygen atoms), which is broken down from rocks in the earth.

Soils have plenty of nitrogen and phosphorus in them. But many do not. Gardening fertilizer contains nitrogen and phosphorus in forms that plant roots can easily slurp up. With all the extra nutrients, the fertilizer ads say, plants will grow bigger and faster.

Explainer: How photosynthesis works

“If [plants] are given a lot of light and nitrogen, they might increase chlorophyll and photosynthesis,” Savage says. That might mean the plants end up with more leaves. With more leaves, she notes, they’ll have more sugar. Those sugars can be made into more plant materials. With fertilizer, Savage explains, plants should get bigger, because they’ll make more sugar.

The question is whether the fertilized plants will have bigger roots, bigger leaves or both. “Will they focus on growing above or below ground?” she asks.

That’s a hypothesis I can test. My hypothesis is that fertilized plants will be bigger than those that are not fertilized.

You don’t need too much stuff for this experiment. A good notebook, small pots, radish seeds, organic potting soil, fertilizer and a spot with plenty of sun. B. Brookshire

Grow radish, grow

I bought several packs of seeds, 24 small plastic seed pots, plant fertilizer and potting soil. I made sure the soil didn’t contain added fertilizer.

I wanted something that I could grow quickly, that wouldn’t take up a lot of space and that wouldn’t get too big. I ran this experiment in early fall in Maryland. So I knew I needed a plant that could grow when it’s cool. I picked radishes, which grow well in the early fall or spring. Some varieties can grow a full radish in only 21 days.

  1. Radish seeds are VERY tiny. Keep track to make sure you don’t lose any. B. Brookshire
  2. Make a small hole about the depth of your fingertip, push in the seed and cover (left). You also want to keep your plants together, so they get the same amount of sun and rain. Label them all so you know which is fertilized and which is not. B. Brookshire

I kept 12 of the pots and one pack of seeds for myself. I gave the other 12 pots and the other packet of seeds — along with some fertilizer and soil — to my editor, Sarah Zielinski. This was to provide an additional control for location. After all, what if my yard just happens to be much better for growing plants? What if it’s worse? By splitting the plants up between my yard and Sarah’s, I hoped to make sure that any difference with the plants came from the fertilizer.

Sarah and I planted our seeds. Sometimes, seeds don’t sprout. So we carefully planted four evenly spaced seeds in each pot. Six of my pots (and six of Sarah’s) served as controls — pots that would not get fertilizer. Our other six were treated with fertilizer. For each of us, this added up to 24 control seeds, and 24 seeds that would get fertilizer.

It’s important to read and follow the instructions for the type of fertilizer you use. (Mine required mixing a tiny capful of liquid with five gallons of water.) Too much can cause fertilizer burn, where plants brown or even die. That’s because the nitrogen in fertilizer mix is in the form of a salt called ammonium nitrate. Such salts in the soil can cause water to leave the plant and head toward the salty soil — a process called osmosis. This can make the plant dry out and look burned.

We watered all the plants equally with clean water every other day (unless it rained). Once a week, we applied fertilizer to half the pots. We also took pictures every day, so we could see the plants change over time.

As I expected, many of our seeds didn’t sprout. In fact, only about a fourth of mine sprouted. Sarah has a greener thumb. She successfully grew half of hers.

Radishing results

The radishes were still too small to weigh, so we measured the root and leaf length. B. Brookshire

Day 21 was the moment of truth! Sarah and I pulled out the radishes, weighed them and measured the leaves and roots.

I pulled out my first radish — and was pretty disappointed.

While these plants can mature in 21 days, that doesn’t mean they always do. Our radishes were pretty puny. But that’s not a bad thing. After all, if all the radishes had grown until they couldn’t get any bigger, it might be harder to see any differences from the fertilizer.

Unfortunately, the radishes were so small that they weighed less than one gram. Most home kitchen scales don’t measure masses that small. Sarah and I were stuck measuring the length of the roots and leaves to see if there was any difference.

We started by counting the leaves on each plant. Together, we grew a total of 30 plants that received no fertilizer. These control plants had an average of 4.1 leaves. We also grew 24 plants in our fertilized pots. These had an average of 5.3 leaves. It seems the fertilized plants had more leaves than control plants.

The control radish (left) looks a good bit smaller than the fertilized one on the right. But does the difference matter? Always grow more than two radishes, enough to do statistics. B. Brookshire

But that doesn’t mean the difference was due to the fertilizer. To find that out, I need to run statistics — tests I can use to interpret my data. In this case, we have two groups — fertilized and control. I used a t test, which can be used to compare two groups to each other. There are lots of sites online that will let you copy and paste in your data. I used this one from GraphPad.

A t test gives you a p value. A p value is a measure of the probability that just by chance I would see a variation between the groups as big as the one I measured. Usually, it’s expressed as a decimal, such as 0.05. That would be a five percent likelihood that I would get a difference as big or bigger than the one I saw if there was no real difference between the groups. Scientists often consider p values smaller than 0.05 to be meaningful — what they call statistically significant.

In this case, the p value between the fertilized and control leaves was 0.0001, or 0.01 percent. That difference is statistically significant. But that doesn’t tell you if the difference between the two is a big one. A difference can be very small and still be statistically significant. To find out if I have a big difference, I need to run a test called a Cohen’s d. You can also run that for free online. I used the calculator here.

This is a table that shows the results of my statistics. The top two rows are the average number of leaves, average root length and average leaf length for control (row one) and fertilized plants (row two). The p value from the t tests comparing the numbers is row three, and the Cohen’s d is row four. You can see that all of my results were statistically significant. But only the leave count and leaf length differences were large.

For the Cohen’s d calculation, I need a number called the standard deviation. This is the amount by which each set of data differs from the mean (or average). To find that, I went to my data in Microsoft Excel, typed in the function “= STDEV” and highlighted my data set. I plugged into my calculator the mean, the standard deviation and the number of plants in each group.

My Cohen’s d was 1.3. Scientists usually consider any number over 0.8 to be a large difference. So it appears that our fertilized plants had more leaves than our non-fertilized controls, and that the fertilizer made a big difference in the number of leaves.

We also measured the length of the leaves and the length of the roots. I’ve included the p values for each one and the Cohen’s d in the table below. Fertilized plants had longer roots, but that difference was not big. They also had longer leaves, and here the difference was again large.

  1. Fertilized plants (right, yellow) had more leaves after three weeks than the control plants (left, blue). B. Brookshire
  2. Fertilized plants (right, yellow) had longer roots after three weeks than the control plants (left, blue). But the difference was pretty small. B. Brookshire
  3. Fertilized plants (right, yellow) had longer leaves after three weeks than the control plants (left, blue). This difference was larger than the difference seen in the root lengths. B. Brookshire

I started with a hypothesis that fertilized plants will be bigger than those that are not fertilized. Well, the fertilized plants had more leaves, and their leaves were longer. The roots were longer too, though the difference wasn’t very large. Overall, it appears that fertilizer does make radishes grow bigger than they might otherwise.

Of course, every experiment has limitations — things that could have gone better. For example, why did so few of my radishes sprout? I think perhaps if I did it again, I would place my pots in a sunnier spot. I also pulled the plants when they were still small. In another experiment, I would give them more time to grow. After all, we got larger radish greens. It’s possible that with more leaves and more time in the sun — and thus more photosynthesis — we’d end up with larger radishes.

There are lots of other things to try. I could try different “doses” of fertilizer. I could also try different types of radish. Maybe some respond better to fertilizer than others. There’s lots of science that can be done with some dirt and a few seeds.

Materials

  • Miracle Gro ($7.48)
  • Organic potting soil (Bumper Crop, $32)
  • Radish seeds (Cherry Belle, 21-day, $1.95/packet)
  • Seedling pots (.50 each)
  • Measuring cups ($7.46)
  • Nitrile or latex gloves ($4.24)
  • Small digital scale ($11.85)

Power Words

atom: The basic unit of a chemical element. Atoms are made up of a dense nucleus that contains positively charged protons and uncharged neutrons. The nucleus is orbited by a cloud of negatively charged electrons.

ATP: Short for adenosine triphosphate. Cells make this molecule to power almost all of their activities. Cells use oxygen and simple sugars to create this molecule, the main source of their energy. The small structures in cells that carry out this energy-storing process are known as mitochondria. Like a battery, ATP stores a bit of usable energy. Once the cell uses it up, mitochondria must recharge the cell by making more ATP using energy harvested from the cell’s nutrients.

average: (in science) A term for the arithmetic mean, which is the sum of a group of numbers that is then divided by the size of the group.

carbon: The chemical element having the atomic number 6. It is the physical basis of all life on Earth. Carbon exists freely as graphite and diamond. It is an important part of coal, limestone and petroleum, and is capable of self-bonding, chemically, to form an enormous number of chemically, biologically and commercially important molecules. (in climate studies) The term carbon sometimes will be used almost interchangeably with carbon dioxide to connote the potential impacts that some action, product, policy or process may have on long-term atmospheric warming.

carbon dioxide: (or CO2) A colorless, odorless gas produced by all animals when the oxygen they inhale reacts with the carbon-rich foods that they’ve eaten. Carbon dioxide also is released when organic matter burns (including fossil fuels like oil or gas). Carbon dioxide acts as a greenhouse gas, trapping heat in Earth’s atmosphere. Plants convert carbon dioxide into oxygen during photosynthesis, the process they use to make their own food.

cell: The smallest structural and functional unit of an organism. Typically too small to see with the unaided eye, it consists of a watery fluid surrounded by a membrane or wall. Depending on their size, animals are made of anywhere from thousands to trillions of cells. Most organisms, such as yeasts, molds, bacteria and some algae, are composed of only one cell.

chemical: A substance formed from two or more atoms that unite (bond) in a fixed proportion and structure. For example, water is a chemical made when two hydrogen atoms bond to one oxygen atom. Its chemical formula is H2O. Chemical also can be an adjective to describe properties of materials that are the result of various reactions between different compounds.

chemistry: The field of science that deals with the composition, structure and properties of substances and how they interact. Scientists use this knowledge to study unfamiliar substances, to reproduce large quantities of useful substances or to design and create new and useful substances. (about compounds) Chemistry also is used as a term to refer to the recipe of a compound, the way it’s produced or some of its properties. People who work in this field are known as chemists. (in social science) A term for the ability of people to cooperate, get along and enjoy each other’s company.

control: (n.) A part of an experiment where there is no change from normal conditions. The control is essential to scientific experiments. It shows that any new effect is likely due only to the part of the test that a researcher has altered. For example, if scientists were testing different types of fertilizer in a garden, they would want one section of it to remain unfertilized, as the control. Its area would show how plants in this garden grow under normal conditions. And that gives scientists something against which they can compare their experimental data. (v.) To include some unchanged or unaffected conditions in an experiment so their results could later be contrasted with those from where changes had been made.

crop: (in agriculture) A type of plant grown intentionally grown and nurtured by farmers, such as corn, coffee or tomatoes. Or the term could apply to the part of the plant harvested and sold by farmers.

data: Facts and/or statistics collected together for analysis but not necessarily organized in a way that gives them meaning. For digital information (the type stored by computers), those data typically are numbers stored in a binary code, portrayed as strings of zeros and ones.

DNA: (short for deoxyribonucleic acid) A long, double-stranded and spiral-shaped molecule inside most living cells that carries genetic instructions. It is built on a backbone of phosphorus, oxygen, and carbon atoms. In all living things, from plants and animals to microbes, these instructions tell cells which molecules to make.

element: A building block of some larger structure. (in chemistry) Each of more than one hundred substances for which the smallest unit of each is a single atom. Examples include hydrogen, oxygen, carbon, lithium and uranium.

fertilizer: Nitrogen, phosphorus and other plant nutrients added to soil, water or foliage to boost crop growth or to replenish nutrients that were lost earlier as they were used by plant roots or leaves.

function: (in math) A relationship between two or more variables in which one variable (the dependent one) is exactly determined by the value of the other variables.

hypothesis: (v. hypothesize) A proposed explanation for a phenomenon. In science, a hypothesis is an idea that must be rigorously tested before it is accepted or rejected.

mass: A number that shows how much an object resists speeding up and slowing down — basically a measure of how much matter that object is made from.

molecule: An electrically neutral group of atoms that represents the smallest possible amount of a chemical compound. Molecules can be made of single types of atoms or of different types. For example, the oxygen in the air is made of two oxygen atoms (O2), but water is made of two hydrogen atoms and one oxygen atom (H2O).

nitrate: An ion formed by the combination of a nitrogen atom bound to three oxygen atoms. The term is also used as a general name for any of various related compounds formed by the combination of such atoms.

nitrogen: A colorless, odorless and nonreactive gaseous element that forms about 78 percent of Earth's atmosphere. Its scientific symbol is N. Nitrogen is released in the form of nitrogen oxides as fossil fuels burn. It comes in two stable forms. Both have 14 protons in the nucleus. But one has 14 neutrons in that nucleus the other has 15. For that difference, they are known, respectively, as nitrogen-14 and nitrogen-15 (or 14 N and 15 N).

nutrient: A vitamin, mineral, fat, carbohydrate or protein that a plant, animal or other organism requires as part of its food in order to survive.

organic: (in chemistry) An adjective that indicates something is carbon-containing also a term that relates to the basic chemicals that make up living organisms. (in agriculture) Farm products grown without the use of non-natural and potentially toxic chemicals, such as pesticides.

osmosis: The movement of certain molecules within a solution across a membrane. The movement is always from the solution where the concentration of some chemical is higher to the solution where the concentration of that chemical is lower. This movement tends to continue until concentrations on each side of the membrane are the same.

oxygen: A gas that makes up about 21 percent of Earth's atmosphere. All animals and many microorganisms need oxygen to fuel their growth (and metabolism).

p: value (in research and statistics) This is the probability of seeing a difference as big or bigger than the one observed if there is no effect of the variable being tested. Scientists generally conclude that a p value of less than five percent (written 0.05) is statistically significant, or unlikely to occur due to some factor other than the one tested.

phosphate: A chemical containing one atom of phosphorus and four atoms of oxygen. It is a component of bones, hard white tooth enamel, and some minerals such as apatite.

phosphorus: A highly reactive, nonmetallic element occurring naturally in phosphates. Its scientific symbol is P. It is an important part of many chemicals and structures that are found in cells, such as membranes, and DNA.

plastic: Any of a series of materials that are easily deformable or synthetic materials that have been made from polymers (long strings of some building-block molecule) that tend to be lightweight, inexpensive and resistant to degradation.

potassium: A chemical element that occurs as a soft, silver-colored metal. Highly reactive, it burns on contact with air or water with a violet flame. It is found not only in ocean water (including as part of sea salt) but also in many minerals.

probability: A mathematical calculation or assessment (essentially the chance) of how likely something is to occur.

protein: A compound made from one or more long chains of amino acids. Proteins are an essential part of all living organisms. They form the basis of living cells, muscle and tissues they also do the work inside of cells. Among the better-known, stand-alone proteins are the hemoglobin (in blood) and the antibodies (also in blood) that attempt to fight infections. Medicines frequently work by latching onto proteins.

salt: A compound made by combining an acid with a base (in a reaction that also creates water). The ocean contains many different salts — collectively called “sea salt.” Common table salt is a made of sodium and chlorine.

seedling: The initial plant that sprouts leaves and roots after emerging from a seed.

standard deviation: (in statistics) The amount that each a set of data varies from the mean.

statistics: The practice or science of collecting and analyzing numerical data in large quantities and interpreting their meaning. Much of this work involves reducing errors that might be attributable to random variation. A professional who works in this field is called a statistician.

sun: The star at the center of Earth’s solar system. It is about 27,000 light-years from the center of the Milky Way galaxy. Also a term for any sunlike star.

Editor's Note:

This story was updated on December 9, 2020 to correct numbers displayed in the table.

About Bethany Brookshire

Bethany Brookshire was a longtime staff writer at Science News for Students. She has a Ph.D. in physiology and pharmacology and likes to write about neuroscience, biology, climate and more. She thinks Porgs are an invasive species.

Classroom Resources for This Article Learn more

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Mr G’s Environmental Systems

Organisms that use inorganic sources of energy, and particularly plants are the base unit of stored energy in any ecosystem. Light energy is converted to chemical energy by photosynthesis within the cells of plants

Because all the energy fixed by plants is converted to sugars it is in theory possible to calculate a plant’s energy uptake by measuring the amount of sugar produced. This is Gross Primary Production (GPP), because it occurs in the primary producers, an abstract that is difficult to measure. More useful is the measure of Net Primary Production (NPP).

An ecosystems NPP is the rate at which plants accumulate dry mass, usually measured in kg,m -2 ,yr -1 , or as the energy value gained per unit time kJ,m -2 ,yr -1 . This store of energy is potential food for consumers within the ecosystem.

NPP represents the difference between the rate at which plants photosynthesize (GPP) and the rate, which they respire (R). This is because the glucose produced in photosynthesis has two main fates. Some provides for growth, maintenance and reproduction with energy being lost as heat during processes such as respiration. The remainder is deposited in and around cells represents the stored dry mass (NPP=GPP-R).

The accumulation of dry mass is more usually termed biomass, and provides a useful measure of the production and use of resources.

Primary production is the foundation of all metabolic processes in an ecosystem, and the distribution of production has a key part in determining the structure of an ecosystem. Biological communities include more than just plants, they also include herbivores, carnivores and detritivores.

Production also occurs in animals as Secondary Production. Importantly though animals do not use all the biomass they consume. Some passes through to become feces. Gross production in animals equals the amount of biomass or energy assimilated or biomass eaten less feces.

As with plants some of the energy assimilated by animals is used to drive cellular processes via respiration the remainder is available to be laid down as new biomass. This is Net Secondary Production. Net secondary productivity (NSP ) = food eaten - faeces - respiration energy


Ask an Expert: What happens to a plant when you put it in a jar?

My 7 year old daughter has a science project due on March 1, 2008 for her first grade class. The experiment is "what happens to a plant when you put it in a jar?" We are required to use the scientific method and will present to the class in early March.

Our hypothesis is that the plant will die. We have two plants, one in the jar and one outside of the jar. Both are on the kitchen counter (I've included a link to the photo of the plants at the bottom of this post). We watered both plants prior to putting the one in the jar. Our thought is that the plant in the jar needs to be left alone for the duration (since it is in the jar, technically, we should not be opening the lid to water and such). We believe that eventually, the lack of nutrients (i.e. water, CO2) will kill the plant in the jar, while the plant outside the jar will thrive because it will be watered and will get CO2.

I am not concerned if our hypothesis is right, because that is the fun of the experiment (learning). However, I do have some questions:

1 - Are we on the right track?
2 - What other types of research should we/could we be doing?
3 - Am I correct to think that we should not be opening the lid on the jar to feed the plant?

Any insight/thoughts would be greatly appreciated.

Re: What happens to a plant when you put it in a jar?

Post by donnahardy2 » Sun Jan 20, 2008 8:40 am

You are absolutely on the right track when you mention fun and learning in doing the project. At this age, I would encourage your daughter to think about and enjoy her project, and do as much as she is able to do.

Plants need carbon dioxide to make sugars, and they produce carbon dioxide during respiration, so I don't know if your plant will run out of CO2. Plants will die if they get too dry, but if the lid is airtight, then the water vapor should not be able to escape from the jar. If the lid is not airtight, and the plant does start to dry out, then you could have your daughter water the plant inside the jar. In science experiments, you want to maintain all conditions identical, except for one variable, and so keeping the soil at the same moisture level would be acceptable. Then any differences in the experiment would be due to the environment inside the jar. But, it's your daughter's experiment, so you should let her decide.

Your experimental design is good. You have the two plants and you have one variable (plant inside and out of a jar). You might encourage your daughter to think about what differences in the environment a jar would cause. If sun shines directly on the jar, there will be a difference in temperature. There will also be a difference in humidity inside the jar, so you might want to explain about evaporation. Can your daughter think of any other differences?

You should help your daughter plan how she is going to present her results now. Is she going to take pictures every few days? This would be good, as measuring plant height, temperature, or humidity is too complicated for a project at this age.

You should definitely get some books on plants from the library and read them to your daughter, if she can't read them. You can help her become an expert on plants so she can explain her project to others.

Re: What happens to a plant when you put it in a jar?

Post by ozweego94 » Sat Mar 01, 2008 11:42 am

Ok. so the project is over and we are reviewing our results. Trying to keep all conditions the same with the exception of one plant in a jar and one out of a jar, the plant in the jar ended up dying - which matched our hypothesis. My only question is - should that have happened? Although the jar was not air tight, it was a small jar. We did water the plant in the jar a much as the one out of the jar - about 2 or 3 times per week. There definitely appeared to be a greenhouse effect going on in the jar, as we did see condensation form on the inside of the jar top and the temperture certainly felt a bit warmer.

Eventually, though, we had to stop watering the plant in the jar as the soil began to look super-moist - almost like it could not hold any more water. Why did this happen? Is it that the covered jar did not allow for enough evaportation? That is my thought. It's like it drowned itself. Or, was the confinement of the jar just too much and ultimately there was not enough air?

I would appreciate some final thoughts. Attached is what the plants looked like at the conclusion of the experiment.


Watch the video: Measurement of Microbial growthTurbidityDryweightFiltrationGATEICAR-NETCSIRRohit Shankar Mane (January 2022).