plant growth parameters

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 Submitted to-      Submitted By-

 Dr. Rakesh Gupta         Abhishek Yadav

                          Roll. No. – P8903A27

                          Section- P8903
Plant growth parameters
The plant growth parameters are those factors by which we the growth of the plants.
The main parameters are height, girth, number of leaves, leaf chlorophyll, leaf color,
number of leaves, total fresh and dry weights. These parameters tell us about the
growth of plant. So these parameters are discussed below one by one –


Height is a variable of fundamental importance when measuring or describing a
tree or forest. Plants may not be defined as trees unless they have the potential to
reach a minimum defined height ,while forests may be distinguished from
woodlands by the number and potential or actual height of the trees.

In forest mensuration, height is used to describe both the quantity and quality of
tree and stand growth:

      Quantity: height is a variable in tree and stand volume tables or equations
       used to predict volume.
      Quanlity: height at a given age is the basis of site index classification, i.e. it
       reflects the fertility of the site.

There are various expressions of tree height:

      total height
      merchantable height and
      bole height.

Techniques for measuring height may also vary depending on whether the tree is:

      straight or
      leaning and if
      on flat or sloping ground.

Total height
Total height is defined as the vertical distance from the base of the tree to the
uppermost point (tip).
In forests where leaning trees are common, measuring the linear distance (or slope
distance) from base of the tree to tip may be more appropriate. The linear distance
will be greater than the vertical distance. However unless the lean is more than
about 8 degrees, the difference between the two is less than 1% . Measurement
crews and data interpeters must be aware which of these two height definitions is

Measurement of total height may be difficult in dense stands of trees with rounded
crowns, e.g. tropical rain forest, and in young plantation tree crops before thinning
opens up the canopy.

Trees with an excurrent habit have a well defined tip. One can measure total
height more accurately in excurrent species. Most conifers are in this group.

Trees with a deliquescent habit have a poorly defined tip. The heights of trees of
deliquescent habit (i.e. umbrageous species) are much more difficult to measure
accurately. The convention is to sight to the point in the crown which is vertically
above the top of the bole and judged to be at the crown surface.

Many hardwood tree species (e.g. most eucalypts) exhibit an excurrent habit when
young, but change to deliquescent habit when they mature.

Merchantable height
Merchantable height concerns conventionally merchantable material only and is
defined as:
the distance from the base of the tree to the highest point of the main stem where
the diameter is not less than some specified value or where utilisation of the bole is
limited by branching or defect.

Merchantable height is therefore a variable quantity which depends on the
specifications for the merchantable product.
Height measurement
Techniques for measuring tree height may be direct or indirect and essentially
depend on the position or fate of the tree:

      Felled trees - when the tree is on the ground, measurement of the linear
       distance from base to tip or to the merchantable limit is done directly with
       linear tape or graduated pole.
      Standing trees - height can be measured by either direct or indirect methods
       (see below). Indirect methods are most common because the tip or
       merchantable limit is often inaccessible.
A height measuring instrument includes a measuring box having a right-angled
plate; a measuring tape having a scale and a space to record measured data, the
measuring tape wounded onto means for winding within the measuring box when in
a stored mode, being extended an optional length when in an operative mode;
winding means provided within the measuring box and attached to one end portion
of the measuring tape so as to wind the measuring tape by handling from the
outside; means for marking the measured data on the measuring tape, the marking
means being included: a measuring member formed in the shape of a plate and
installed into the measuring box such that the measuring member can prevent to
loosen and slide the measuring tape in an operative mode by a tension on the
measuring tape when in fitting of a length of the measuring tape; a through hole
formed on the measuring member and allowing writing goods to pass through
therein order to record a mark on the measuring tape; a hole punch having a spring
   provided fixedly at the measuring member such that the hole punch mark the
   measured data on the measuring tape; and a step attached to another end portion of
   the measuring tape. Because the height measure is formed in this way, the
   measuring tape recorded the correct measured data of the height can be wound
   onto the shaft within the measuring box easily when the instrument not in measuring
   use, and it is a portable simply. Also, the measuring tape data can be kept as
   memorial goods for life.

   The girth of a tree is much easier to measure than the height, as it is a simple matter
   of stretching a tape round the trunk, and pulling it taut to find the circumference.

The girth of the plant is measured with the help of following instruments-

 A Biltmore stick is a tool used to measure both tree diameter (girth) and height. A
diameter tape is used to get a more accurate measurement of tree diameter.


In this study, harpin protein was applied to the peppers (Capsicum annuum L. var. cvs.
‘Demre’, ‘Yalova Charleston’ and ‘Sari Sivri’) grown under natural conditions. These
plants were subjected to artificial inoculation with Botrytis cinerea, which causes fruit
spoilage in peppers. Changes in vegetative growth, total chlorophyll content in leaves,
leaf colour and percentage of rotten fruits were determined after treatments. The
number of leaves per plant value was quite low in all cultivars and the plant height value
was low only in cv. ‘Sari Sivri’ treated with B. cinerea. Values obtained from vegetative
growth parameters in the plants subjected to harpin protein + B. cinerea treatment were
only higher than B. cinerea treatment. Leaf chlorophyll values exhibited significant
decline in the plants subjected to B. cinerea treatment in all cultivars. However, the
chlorophyll content in the plants subjected to harpin protein + B. cinerea treatment was
low. The colour values obtained from leaves supported the chlorophyll findings. Fruit
spoilage percentages were lower in the fruits picked from the plants of harpin
protein + B. cinerea treatment compared with those picked from the plants only
subjected to B. cinerea treatment.

In this study, two sweet sorghum cultivars (Soave and Sofra) were grown in

nutrient solution containing 0, 30, 60 and 90 mM NaCl to evaluate the effects of salinity

growth parameters (shoot fresh weight, shoot dry weight, leaf area index and relative

chlorophyll) and carbohydrate content (sucrose, glucose and fructose) in plant organs

(leaf, shoot and root). The results showed that as salinity increased, the amount of the

growth parameters in plant parts of both Soave and Sofra decreased. However in all

salinities, Soave had significantly higher shoot fresh weight and leaf area index than

As salinity increased, the amount of sucrose and glucose of both Soave and Sofra

while the amount of fructose increased. However, the sucrose content of Soave was

than Sofra but its glucose and fructose content was lower than Sofra. Since salinity

shoot fresh weight and leaf area index, mainly in the salt sensitive plants. Also, salinity

increases carbohydrate contents especially in the salt tolerance plants. Therefore, it

that Soave is more salt tolerance than Sofra.

IN investigations of plant development it is helpful to be able to represent
diagrammatically the changes which occur in the distribution of dry weight, total nitrogen
or some other variable as the plant grows. It is usually desirable to distinguish at least
four components of the plants of different physiological function, namely, leaf laminae,
stems plus petioles, flowers plus fruits and roots, though data for the last-mentioned are
often lacking. The values of the variable for each of these components should be plotted
against a suitable independent variate which may be, for example, age, node number,
leaf area or total dry weight. If the variable for the four components is plotted as four
ordinary two-dimensional graphs, whether independently from the same base line or
superimposed to build up the total, it is not easy to see whether the ratios of the values
for the different components are constant or changing with growth of the plant. This can
be seen by plotting logarithms of the values of the variable, but then their absolute
magnitudes are obscured.

Advanced selections (families 20010 and 20062) of P. radiata D. Don were exposed to
either 340 or 660 μmol CO2 mol 1 for 2 years to establish if growth responses to high
CO2 would persist during the development of woody tissues. The experiment was
carried out in glasshouses and some of the trees at each CO2 concentration were
subjected to phosphorus deficiency and to periodic drought. CO2 enrichment increased
whole-plant dry matter production irrespective of water availability, but only when
phosphorus supply was adequate. The greatest increase occurred during the
exponential period of growth and appeared to be tied to increased rates of
photosynthesis, which caused accelerated production of leaf area. The increase in
whole-plant dry matter production was similar for both families; however, family 20010
partitioned larger amounts of dry weight to the trunks than family 20062. which favoured
the roots and branches. Wood density was generally increased by elevated CO 2 and for
family 20010 this increase was due to thickening of the tracheid walls. Tracheid length
was similar at both CO2 levels but differed between families. These results suggest that,
as the atmospheric CO2 concentration rises, field-grown P. radiata should produce more
dry weight at sites where phosphorus is not acutely deficient, even where drought limits
growth; however, increases in wood production are likely only for genotypes which
continue to partition at least the same proportion of dry weight to wood in the trunk.
Fresh-weight measurements of roots provide inaccurate estimates of the effects of plant
growth-promoting bacteria on root growth: a critical examination

Four strains of plant growth-promoting bacteria (PGPB), including three strains of Azospirillum
and Pseudomonas fluorescens 313 were used to inoculate seeds of wheat, tomato, pepper, and
cotton. Inoculated seedlings were grown to the two or three-leaf stage. After harvest, seven
different environmental and technical conditions were evaluated to determine the effect of
these conditions on the reproducibility of fresh and dry root weight measurements. Dry root
weight of each sample showed no significant variations (smaller than 1%), despite the
variations in measurement conditions. Root fresh weights varied greatly (usually in the range of
4–10%, but up to 18%), and were significantly affected (P≤0.05) by air temperature and relative
humidity, air currents, different light intensities during extraction of plants from the substrate,
duration of the extraction from soil (depending on the size of the experiment and type of plant
growth substrate), and the type of absorbing paper used to blot excess water from harvested
roots. Measurements by different technicians did not influence fresh or dry weight values. We
conclude that fresh weight determinations are altered by factors independent of the intended
experimental variables and should not be used to evaluate the effect of PGPB on plants.

Weighing Plants: Fresh vs. Dry Weight

      Measuring Fresh Weight: While you can technically measure the fresh weight of plants
       without harming them, the simple act of removing a plant from its growing "medium"
       can cause trauma and affect the ongoing growth rate and thus your experiment.
       Measuring the fresh weight of plants is tricky and should probably be saved as a final
       measure of growth at the end of the experiment. Here is the process for measuring
       fresh weight:

           1. Remove plants from soil and wash off any loose soil.

           2. Blot plants gently with soft paper towel to remove any free surface moisture.

           3. Weigh immediately (plants have a high composition of water, so waiting to
              weigh them may lead to some drying and therefore produce inaccurate data).
      Measuring dry weight: Since plants have a high composition of water and the level of
       water in a plant will depend on the amount of water in it's environment (which is very
       difficult to control), using dry weight as a measure of plant growth tends to be more
       reliable. You can only capture this data once as a final measure at the conclusion of your

            1. Remove the plants from the soil and wash off any loose soil.

            2. Blot the plants removing any free surface moisture.

            3. Dry the plants in an oven set to low heat (100 degrees) overnight.

            4. Let the plants cool in a dry environment (a Ziploc bag will keep moisture out) - in
               a humid environment the plant tissue will take up water. Once the plants have
               cooled weigh them on a scale.

            5. Plants contain mostly water, so make sure you have a scale that goes down to
               milligrams since a dry plant will not weight very much.

Root Mass

Root mass is recommended as a final measurement as the plant must be removed from it's
growing medium in order to capture accurate data. There are quite a few different methods for
measuring root mass depending on the type and structure of the roots

      Grid intersect technique:

            1. Remove the plant from the soil.

            2. If you are working with thin or light roots, you may want to die the roots using
               an acidic stain.

            3. Lay the roots on a grid pattern and count the number of times the roots intersect
               the grid.

      Trace the roots on paper, measure each of the tracings, and calculate root length from
       the tracings.

      Count the number of roots.

      Measure the diameter of the root. This is especially useful for root vegetables such as
       beets, carrots, potatoes, etc. that have a large root.
Root Shoot Ratio

Roots allow a plant to absorb water and nutrients from the surrounding soil, and a healthy root
system is key to a healthy plant. The root:shoot ratio is one measure to help you assess the
overall health of your plants. Your control group of plants will provide you with a "normal"
root:shoot ratio for each of your plant types, any changes from this normal level (either up or
down) would be an indication of a change in the overall health of your plant. It is important to
combine the data from the root:shoot ratio with data from observations to get an accurate
understanding of what is happening with your plants. For example, an increase in root:shoot
ratio could be an indication of a healthier plant, provided the increase came from greater root
size and NOT from a decrease in shoot weight. To measure the root:shoot ratio:

   1. Remove the plants from soil and wash off any loose soil.

   2. Blot the plants removing any free surface moisture.

   3. Dry the plants in an oven set to low heat (100 degrees) overnight.

   4. Let the plants cool in a dry environment (a Ziploc bag will keep moisture out) - in a
      humid environment the tissue will take up water. Once the plants have cooled weigh
      them on a scale.

   5. Separate the root from the top (cut at soil line).

   6. Separately weigh and record the root and top for each plant. (Dry weight for roots/dry
      weight for top of plant = root/shoot ratio)

   7. The root/shoot ratio can be calculated for each treatment.

   8. Plants contain mostly water, so make sure you have a scale that goes down to
      milligrams since a dry plant will not weight very much.


There are many different features of a plant that can be measured through observation to
determine the extent of plant growth/health. The following table describes some of the
measures that you can make and also recommends how frequently you should make these
observations during the course of your experiment.

               Measurement          Procedure                                                 Frequency of

When starting with seeds First Cotyledon   Record the number of days from planting to the            Once
                                           emergence of the first cotyledon ("seed leave(s)" that
                                           are the first to emerge from the ground).

                         Percentage of     Calculate the percentage of seeds that germinated         Once
                         seeds that        under each of the variables in your experiment.

When starting with young Plant height            Measure the height of the main plant from the      Every 2-3 days
plants                                            border of the container to the top of the main
                                                  plant stem.

                                                 Note: you do not want to measure from the top
                                                  of the soil, as the soil may condense with
                                                  watering over time.

                         Number of leaves Counting Leaves: (4)                                Every 2-3 days
                         (indicates a plant's
                         physiological age)    Count and record the number of leaves on each

                                                 Count every visible leaf on the plant, including
                                                  the tips of new leaves just beginning to emerge.

                                                 You may want to place the plant over some
                                                  graph paper to avoid counting errors.

                         Surface area of         Method 1 (5): Trace the leaves on graph paper Every 2-3 days
                         leaves                   and count the squares covered to give you an
                                                  estimate of the surface area for each leaf.
                                                  Repeat this for each leaf on a plant and for each
                                                  plant in your experiment.

                                                 Method 2 (6): Trace out each leaf on paper.
                                                  Make sure to use the same type of paper every
                                                  time AND make sure that the paper is not wet.
                                                  Cut out the leaf tracings and weigh them.
                                                       Weigh the cutouts and divide the total weight
                                                       by the number of leaves to give you the average
                                                       leaf area for each plant. Repeat this for each of
                                                       the plants in your experiment.

                                                      Method 3 (7): Digital image analysis: Using a
                                                       digital camera capture an image of a plant.
                                                       Using special software, analyze the surface area
                                                       of the leaves.

                           Plant color          Record any observations on changes or differences in       Every 2-3 days
                                                plant color.

When you are using         1st Flowering        Record the number of days since initial planting to the    Once
flowering plants these                          first flower.
two measurements serve
as an additional
indication of plant health

                           Number of Flowers Record the number of flowers on each of the plants.           Every 2-3 days
                                             Buds should be included in your flower count.

            An auxanometer (Gr. auxain, "to grow" + metron, "measure") is an apparatus for measuring
            increase or rate of growth in plants.

            In case of an arc-auxanometer (see picture), there is a wire fixed with the plant apex on one
            end and a dead-weight on the other. It passes over a pulley which has a pointer attached to it.
            When the plant's height increases, the pulley rotates and the pointer moves on a circular scale
            to directly give the magnitude of growth.

            Sensitive auxanometers allow measurement of growth as small as a micrometer, which allows
            measurement of growth in response to short-term changes in atmospheric composition.
            Auxanometers are used in the laboratory, the field, and the classroom.
A Modern Tool for Classical Plant Growth Analysis

We present an all-inclusive software tool for dealing with the essential core of mathematical
and statistical calculations in plant growth analysis. The tool calculates up to six of the most
fundamental growth parameters according to a purely ‘classical’ approach across one harvest-
interval. All of the estimates carry standard errors and 95 % confidence limits.

Controlling plant growth via the gibberellin biosynthesis system – I. Growth parameter
alterations in apple seedlings

'York Imperial' apple seedlings (Malus domestica Borkh.) were continuously supplied via the
roots with paclobutrazol [(2RS, 3RS)-1-(4-chlorophenyl)-4,4-dimethyl-2-(1,2,4-triazol-1-
yl)pentan-3-ol)], a triazole GA biosynthesis inhibitor, at 0.68 μM in a nutrient solution. In
comparison to controls, seedlings treated with paclobutrazol for 66 days showed a 91%
reduction in shoot length, a 66% reduction in leaf area but only a 17% reduction in leaf number.
This effect could be reversed by GA3 applied to the foliage at 71.4 μM 0, 19 or 35 days after
paclobutrazol was initially supplied and leaf area values for paclobutrazol-treated seedlings
given both treatments did not differ significantly from controls. Plots of growth data indicate
linearity of shoot longitudinal growth of GA3-treated seedlings. Leaf area increase was non-
linear after GA3 treatment up to approximately 30 days, when the rate dropped. On a per shoot
basis, leaf weight closely followed leaf area but on a per unit area basis, paclobutrazol-treated
leaves were heavier than controls; GA3 applications temporarily reversed this trend.

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