BIOLOGY 468/568: Principles and Applications of Electron Microscopy

Species Identification
Kingdom: Plantae
Division: Spermatophyta
Subdivision: Angiospermae
Class: Monocotyledoneae
Order: Graminale
Family: Poaceae
Genus: Arundo
Species: donax

Key Words: mitochondria, roots, Arundo donax

 

INTRODUCTION

Marshes, estuaries and riparian habitats are important as breading grounds for various native and non-native species. These three ecotones are greatly affected by water availability and salt water influx, which in turn determines the types of plant and animal species that can be found in the surrounding waters. On the East Coast, the most dominant marsh plant species are the Spartina grasses. These plants are able to live in a constant salt environment with fairly strong current, and take in salt water through the root membranes that are able to exclude the salt. Any salt that does get in can be removed by depositing it into vacuoles which then take the salt to salt glands in their leaves for extrusion. These salt levels must be balanced or the plant cells will die through loss of water due to osmosis. Spartina alterniflora is bushy and can grow to be ten feet tall; thus causing little light to reach the soil beneath it, wind to be dissipated through its stems, and humidity and temperature to increase in surrounding areas (14). It has been suggested to me that Arundo donax is similar to Spartina alterniflora (17).
Arundo donax or the "giant reed" is a member of the grass family (Poaceae). It can grow up to 25 feet tall, and propagates through rhizomes or fallen branches, with their seed rarely being fertile. Because of this, any stems that are washed down stream of the original plant can take root and grow, making this weed very difficult to eradicate. This ease in propagation is a problem for several reasons. For one, A. donax changes flood-defined riparian forests into fire-defined A. donax habitat because it is highly flammable, and uses three times more water to grow than native plants. Therefore, when a fire starts, it spreads quickly, is very hot, kills everything, and enables A. donax to spread and root faster than native species which must grow from seeds. This effectively eliminates the cottonwood/willow forests along with their defining shrubs that are needed by our native wildlife as habitat for survival; there is no evidence that A. donax is used as either food or habitat for our native species. The problem is compounded by the fact that unlike native species, A. donax provides no shade to the surrounding environment. This causes the temperature of the soil and water to increase, which eliminates aquatic diversity of native breeding fishes, and increases the water’s pH by supplying light for algae to grow. This in turn causes higher levels of ionized ammonia to exist, which is toxic to any native plant that could possibly be left (4). Finally, A. donax has been seen to be able to propagate in salinities higher than what is tolerable for native species. It is believed that salt intake/exclusion will require more cellular energy, resulting in a greater number of mitochondria. Therefore, transmission electron microscopy was used to compare the numbers of mitochondria found in A. donax roots grown at salinities of both 0/00 and 15/00.

 

MATERIALS AND METHODS

Root Growth (all done by George Peck):
Ten prostrate stems of A. donax were collected at the Palos Verdes’ Land Conservancy in Palos Verdes, California, USA on March 3, 1997; the San Gabriel River, Long Beach, California, USA on March 11, 1997; and the Los Angeles River, Los Angeles, California, USA on March 20, 1997. These stems were then cut into 10 cm lengths, divided into three treatments (0/00, 15/00, and 30/00 salinity), and grown hydroponically in tap water/salt treatment starting on March 21, 1997. The roots were grown until April 14, 1997, with growth solutions being replenished as necessary. Roots were only collected from the 0/00 and 15/00 salinity treatments from Palos Verdes and the San Gabriel River.


Figure 1: Shoot cuttings of Arundo donax planted in soil for another experiment done by George Peck. The cuttings used in this experiment were of the same size and design, but were grown hydroponically.

 
Figure 2: The hydroponics set up of Arundo donax shoot cuttings. This is a view of the underside, showing root growth.

 
Figure 3: Root growth comparison of 0/00 (top) and 15/00 (bottom) on Arundo donax shoots that had been grown for approximately three weeks.

TEM:
          Four roots were cut from the Palos Verdes and San Gabriel River stems for both the 0/00 and 15/00 salinity treatments, resulting in a total of eight roots per salinity treatment. Each root was cut approximately four centimeters up from the root tip. The cutting was done under water, in the greenhouse, to avoid air from entering the roots. The roots were then cut under 5% gluteraldhyde, in the lab, with the last two centimeters of each root being discarded just in case the ends had been penetrated by air. These samples were then fixed in 5% glutaraldhyde then osmium tetroxide and dehydrated with an alcohol series of 30%, 50%, 70%, 95%, and 100% ethanol. Half of each sample was removed and saved for SEM preparation. The remaining samples were infiltrated with propylene oxide and Spurr resin; and embedded in 100% Spurr resin. Thin sections were obtained on a thermal advance microtome, stained with uranyl acetate and lead citrate, and viewed with the Joel: JEM - 1200 EX II Electron Microscope at 80kV.

 SEM:

The samples that had been removed after the ethanol series were cryofractured using liquid Nitrogen and a scalpel. They were then returned to 100% ethanol, critically point dried using liquid Carbon Dioxide, and sputter coated with gold/palladium. The samples were viewed on a Joel: JSM - T200 Scanning Microscope at 25kV.

 Processing data:
Photographs were taken with both microscopes and prints were made from the negatives using an Omega Pro Lab 4x5 Enlarger. The prints were scanned with a Hewlett Packard Scan Jet IICX at a resolution of 200 pixels, and contrasts were increased with Aldus PhotoStyler 2.0. Informational tags were added using Photoshop.  The number of mitochondria within the 0/00 and 15/00 treatments were counted on the TEM. Twelve cells were arbitrarily selected from each grid used, and the resulting number of mitochondria were analyzed using Minitab’s Mann-Whitney Test.

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RESULTS

The roots obtained through hydroponic growth were extremely different in size and morphology. The roots of the 0/00 salinity treatment were up to approximately 14 cm long, and contained many lateral roots in the upper 8 cm (Figure 3). In contrast, the 15/00 salinity treatment’s roots were only up to approximately 6 cm long and contained no lateral roots. This suggests that the 15/00 salinity treatment induced considerable stress on the growing roots, which might produce different metabolic needs for the plants.

 

 

No significant difference was found between salinity treatments on the counted mitochondrial number (p= 0.1939).

 

The results of the SEM showed that the root tips of both salinity treatments were actively loosing cells in the root cap vicinity (Figure 4). This is as expected for actively growing roots. Figure 4 also shows what appear to be mycorrhiza on the root tip. If these are in fact mycorrhiza, then the A. donax roots are receiving help in fixing nitrogen, which could increase its ability to out compete other species. The SEM also showed that both salinity treatments contained root hairs (Figure 5). This was unexpected because only the 0/00 roots contained lateral roots after the three week growing period. Since both lateral roots and root hairs help increase water and nutrient uptake, it was assumed that the 15/00 treatment would not contain root hairs. Lastly, the SEM showed that the possibility of achieving a clean cell fracture is extremely difficult when using the liquid Nitrogen Cryofracture Technique on plant tissue. Of the over 30 fractions I made, only one was between cell wall membranes (Figure 6).


Figure 4: The root tip of 15/00 salinity treatment of Arundo donax. The appearance of the root tip of the 0/00 treatment was the same. (Rt = root tip; Rc = root cap cells that are dying; arrow = mycorrhiza or remnant mucigel).

 
Figure 5: Root hairs on a 0/00 salinity root of Arundo donax. The appearance of root hairs was the same on the 15/00 salinity roots. (Rh = root hair).

 
Figure 6: Whole cell fractures showing some of the internal structure of Arundo donax root cells. (N = nucleus; Cw = cell wall; mt = microtubules).

 

The TEM showed that starch grains are accumulating in amyoplasts in both salinity treatment (Figure 15), that vacuoles are filled with a granular material (Figures 12, 13, 14), and that there is either mitochondria in two different conformational states, or that there are mitochondria and microbodies or proplastids in both salinity types (Figures 7, 8, 9, 10, 11). Either situation holds possible significant implications.


Figure 7: A "dark" and "light" mitochondria in a 0/00 salinity root cell. (mag. = 50k; M = mitochondria; C = cristae; cr = possible cristae or endoplasmic reticulum; arrow = possible mitochondria, plastid, or microbody).

 
Figure 8: A cell showing a mixture of "dark" and "light" mitochondria after 15/00 salinity treatment. (mag. = 25k; M = "dark" mitochondria; arrow = "light" mitochondria, plastid, or microbody).


Figure 9: A "dark" mitochondria from the 15/00 salinity treatment. The magnified image of "dark" mitochondria from the 0/00 salinity treatment is the same in appearance. (mag. = 80k; M = mitochondria; C = cristae).


Figure 10: A "light" mitochondria from the 15/00 salinity treatment. The number of possible cristae seen in each salinity's "light" mitochondria was variable, but the overall appearance was the same. (mag. 80k; cr = possible cristae; arrow = possible mitochondria, plastid, or microbody).


Figure 11: A splitting organelle in a 0/00 salinity root cell of Arundo donax. This could either be a splitting mitochondria, microbody, or plastid. (mag. = 50k; Mp = parent organelle; mp = daughter organelles).

 
Figure 12: An entire 0/00 salinity cell. (mag. = 3k; M = "dark" mitochondria; V = vacuole with granular material; Cw = cell wall; N = nucleus; arrow = "light' mitochondria).


Figure 13: A whole root cell from the 15/00 salinity treatment. (mag. = 3k; M = "dark" mitochondria; V = vacuole containing granular material; Cw = cell wall; N = nucleus; arrow = "light" mitochondria).


Figure 14: An entire 0/00 salinity treatment root cell showing amyoplasts. The 15/00 salinity treatment root cells contained amyoplasts as well. (mag. = 4k; M = "dark" mitochondria; V = vacuole containing granular material; Cw = cell wall; S = amyoplast filled with starch).


Figure 15: A magnified view of an amyoplast. (mag. = 30k; M ="dark" mitochondria; S = amyoplast filled with starch).
 

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DISCUSSION

Salt stress in plants is similar to drought stress. In both cases, osmotic differences between the plant and its environment result in water loss. Plants growing in saline environments have adopted many ways to deal with this stress. These include collecting salt in vacuoles, extruding salt form leaves, and excluding salt from water uptake at the root/water interface. The TEM showed that the vacuoles of A. donax roots contained a granular material (Figures 12, 13). This material may either be starch or salt. Since amyoplasts were seen to be present in other root cells (Figure 14, 15)- (3), it is more likely that these grains are salt. Only future research on their chemical make up will be able to determine if this assumption is correct. Another adaptation to surviving in high salinities might be a symbiotic relationship of the roots with nitrogen fixing bacteria or fungi. This association would bypass the problem of bring in/excluding ammonium ions in competition with sodium ions. The SEM showed tiny strand -like growths between root cap cells (Figure 4). They are similar in appearance to mycorrhiza described by Juniper and Jeffree. However, they could also be mucigel remnants from the root cap. Once again, further study of these structures is necessary before a conclusion can be made.
The non-significant results of the Mann-Whitney test imply that A. donax roots under salt stress does not have increased energy requirements for water uptake. This result is explainable by the observation of two mitochondrial types existing within a single A. donax root cell. These have been classified as "light" and "dark" mitochondria. It is possible that it is the ratio of these two that is important, rather than the final number of mitochondria that are pressent within the cells. This is because mitochondria in plants have a variety of sizes, shapes, and configurations depending on their metabolic state at the time of fixation. The different sizes and shapes of mitochondria may be due to different mitochondrial populations within the cell, but is more likely a result of space constraints within the cytoplasm (7). This is supported by the observation that mitochondria isolated from cells are round. Also, mitochondria in vivo have been seen to move, branch, split, and re-form (7), and to reproduce like bacteria (12)-(Figure 11). It has further been shown that different conformational states are a result of the age, type, metabolic, and aeration states of the tissue they are located in (1,6,8,15), and that their appearance can be altered by the plane of orientation they were cut through (16). These conformational states are interconvertable, with two or more conformations being able to co-exist in their combined medium (1,6,8,15). However, it has also been shown by Avers & Tkal (1), that two mitochodrial states can exist in vivo within one cell. Similarly, it has been shown that microbodies (single membrane organelles containing granular material) and proplastids (double membrane organelles also containing granular material) are found in the root cells of plants, and can contain cellular invaginations that appear similar to mitochondria cristae under the TEM (9,11,13).
Based on the above information, the correct identification of mitochondria within A. donax root cells was difficult. The identification of “dark” mitochondria is most likely correct because they consist of many darkly stained cristae that are comparable to traditional mitochondrial micrographs (1,6,8,15). These "dark" mitochondria were seen in both salinity treatments (Figure 7, 8). The identification of the “light” mitochondria, so named because of they contained fewer, lighter stained cristae, is confused on many ambiguities.
For one, as shown by Douce (7), mitochondria in different osmotic mediums result in different sizes and condensed forms. Specifically, in an iso-osmotic medium mitochondria appear to be in a normal configuration; while a high-osmolarity medium causes the cristae to become super condensed and the cell to shrink. Conversely, a low-osmolarity medium causes the cristae and cell to swell to the point that the cristae can almost disappear, and the cell can rupture. A medium such as salt water, containing more solutes than the mitochondria would result in the condensed conformation. Since the level of solutes within a cell is fairly consistent, not all of the mitochondria had this swollen conformation, and the two types were sometimes found adjacent to each other (Figure 7, 8), it is unlikely that the swollen conformation of the “light” mitochondria or the condensed appearance of the "dark" mitochondria is a result of differing osmoregularities.
The correct identification of the “light" mitochondria as such was also difficult because of the findings by Parsons et. al (13). In their experiment, they used different techniques to isolate mitochondria and plastids from plant cells. Their micrographs show mitochondria and plastids that look like my “light" mitochondria. Similarly, Chaffey's (5) micrographs show mitochondria and microbodies that look similar to my “light mitochondria”. Finally, Beezley et al ‘s (2) micrographs show a mitochondria and microbodies that were differentially stained. The unstained microbody looks similar to my “light" mitochondria. This leads me to believe that my “light" mitochondria could be either a microbody or a plastid.
However, as mentioned previously, their are several studies that show mitochondria can exist in different conformational states at the same time once separated from their cells. For example, Deamer et. al (6) showed that adding sodium phosphate, which energizes ion transport into the cell, causes conformational changes in rat liver mitochondria. Specifically, the mitochondria were seen to swell and then return to their original state. This suggests that my “light" mitochondria are in fact mitochondria and not microbodies or plastids. However, the fact that rat liver was used, and that the cells were in vitro makes me reluctant to make this conclusion. Similarly, Hackenbrock (8) used in vitro mouse liver mitochondria to show that changes in metabolic states induced by the addition of sucrose and measured by oxygen evolution, resulted in five different conformational states. Theses states were interconvertable. Once again, this leads me to believe that my “light" mitochondria could be mitochondria in different metabolic states but, since mouse liver was used and not plant cells, the comparison to my study is not strong.
Conversely, two studies do exist that present stronger support for my belief that my “light" mitochondria are in fact mitochondria and not some other type of organelle. The first study was done by Watson et.al (15) on the effects of a sorbitol gradient and aeration states on yeast mitochondria. They found that conformational state changes occur in vivo and that these states were affected by differing amount of oxygen and lipid supplements. They also found that these differing treatments resulted in different metabolic activity of the mitochondria. The second, and most relevant study, was done by Avers and Tkal (1) on the mistematic cells of timothy grass seedling roots. They found that inducing different enzyme activity within the intact cells resulted in two different forms of mitochondria within the root cells. These two forms were found to be either enzymatically active or inactive. This implies that there are different molecular and/or submolecular events occurring within one cell. Based solely on the information obtained in these two studies, it appears that the “light" mitochondria I found in A. donax roots are in truth mitochondria.
In summary, the study of Arundo donax roots and root cells through the use of SEM showed that the roots had the same general morphology (sloughing cells and root hairs) in the area sampled, and that mycorrhiza may be present on the root cap. The TEM showed that there was no significant difference in the number of mitochondria seen, but this could be due to the observation that two supposedly different types of mitochondria were found. It is possible that if these two types are both in fact mitochondria, and that their ratio is more important than their number. This seems likely when we consider that Avers and Tkal (1) found both enzymatically active and inactive mitochondria within one plant cell.
In conclusion, I am wary of stating that there are two types of mitochondria in the root cells of A. donax because of the similarity in appearance of the “light" mitochondria to both microbodies and plastids, both of which are known to be found in plant root cells. Future research needs to be done using differential staining techniques which can visibly separate mitochondria from plastids and microbodies (2,5). If it is found that they are mitochondria, than the enzyme activity status of the two different mitochondrial types should be determined in a manner similar to that described by Avers and Tkal (1). Finally, if it is found that there is an enzymatic difference, then I believe that Arundo donax and native plant species root's should be compared. The presence or absence of similar results in these plants might offer a further evidence as to why A. donax is so much better at out-competing our native plant species.  

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LITERATURE CITED

1. Avers C. J. and M. M. Tkal. 1962. Intracellular mitochondrial variation in enzyme activity as shown by histochemical studies using light and electron microscopy. American Journal of Botany. (Volume???): 157-162.
2. Beezley B. B., Bruber P. J. and S. E. Frederick. 1976. Cytochemical localization of glycolate dehydrogenase is mitochondria of Chlamydomonas. Plant Physiology. 58: 315-319.
3. Bewley J. D. and M. Black. Physiology and Biochemistry of Seeds: In relation to germination. Springer-Verlag, Berlin. 1978. pp. 30-32.
4. California Exotic Pest Plant Council. Arundo donax Workshop Proceedings. 1993.
5. Chaffey N.J. 1985. Structure and Function in the Grass Ligule: Mitochondria and microbodies in the membranous ligule of Lolium temulentum L., an ultrastructural and cytochemical study. Journal of Experimental Botany. 36(167): 1001-1007.
6. Deamer D. W., Utsumi K. and L. Packer. 1967. Oscillatory States of Mitochondria: III. Ultrastructure of trapped conformational states. Archives of Biochemisrty and Biophysics. 121: 641-651.
7. Douce R. Mitochondria in Higher Plants: Structure, function and biogenesis.Academic Press, INC, Orlando. 1985. pp. 1-35.
8. Hackenbrock C.R. 1966. Ultrastructural Bases for Metabolically Linked Mechanical Activity in Mitochondria: I. Reversible ultrastructural changes with change in metabolic steady state in isolated liver mitochondria. The Journal of Cell Biology. 30: 269-297.
9. Hruban Z. and M. Rechcigl Jr. Microbodies and Related Particles: Morphology, biochemistry, and physiology. Academic Press, New York. 1969. pp. 79-86.
10. Juniper B.E. and C. E. Jeffree. Plant Surfaces. Edward Arnold Publishers, (City???). 1983. pp. 55-75.
11. Kirk J. T. O. and R. A. E. Tilney-Bassett. The Plastids: Their Chemistry, Structure, Growth and Inheritance.W. H. Freeman and Company, London. 1967. pp. 63-72.
12. Osumi M. and N. Sando. 1969. Division of yeast mitochondria in synchronous culture. Journal of Electron Microscopy. 18(1): 47-56.
13. Parsons D. F., Bonner W. D. Jr., and J. G. Verboon. 1965. Electron microscopy of isolated plant mitochondria and plastids using both the thin-section and negative-stainig techniques. Canadian Journal of Botany. 43: 647-655.
14. Teal J. and M. Life and Death of the Salt Marsh. Little, Brown and Company, Boston. 1991. pp 84-101.
15. Watson K., Haslam B., Veitch B. and A. W. Linnane, in: Mitochondrial precursors in anaerobically grown yeast.  Autonomy and Biogenesis of Mitochondria and Chloroplasts. North -Holland Publishng Company, Amsterdam. 1971. pp. 162-174.
16. Wheater P. R., Burkitt H. G., and V. G. Daniels. Functional Histology: A text and colour atlas. Churchill Livingstone, London. 1987. pp. 17-19.
17. Wijte A. Personal communication. May 15, 1997.

ACKNOWLEDGMENTS

I'd like to thank George Peck for graciously donating the roots of his hydroponics experiment, as well as his accompanying photographs. I'd also like to thank Dr. Zed Mason, Dr. Tom Douglas, Dr. Antonia Wijte, Danny Tang, Todd Chapman, and Robert Engen, all of which helped to make this project successful in some way or another. Thanks for your time, expertise, help and patience. 

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Questions? Comments? Then mail me.