- Saprophyte Definition
- Function of a Saprophyte
- What Is A Saprophyte And What Do Saprophytes Feed On
- What is a Saprophyte?
- What Do Saprophytes Feed On?
- Additional Saprophyte Information
- RESULTS AND DISCUSSION
A saprophyte, also referred to as a saprobe or saprotroph, is any organism that feeds and grows on dead organisms. This means that a saprophyte is a decomposer, breaking down complex matter and absorbing the simpler products. Since saprophytes rely on dead plant and animal bodies for food, rather than producing their own as autotrophs do, they are heterotrophs. Keep in mind that although it is still used, saprophyte may be a misleading name, since –phyte means plant. What makes this an issue is that it has been found that no land plants truly feed in the manner that a saprophyte does, but it may seem like it when plants use fungi to acquire nutrients.
Saprophytic nutrition is usually displayed by bacteria and fungi living in moist environments. They decompose organic dead and decaying matter by extracellular digestion, which is the secretion of digestive juices that break down matter around them. In the case of fungi, we find that most are multicellular saprophytes. They grow tubular structures, or hyphae, which are filaments that grow and branch into the dead matter, produce digestive enzymes, and digest away the dead organism. The fungi then absorb the simple substances through their hyphae, which can in time grow into a mycelium, or a mass of hyphae, as seen below.
While extracellular digestion is the means by which most fungi and bacteria acquire their nutrition, bacteria are simpler organisms and do not produce hyphae.
In addition to their preference for humid environments, most saprophytes require oxygen to survive and would die in its absence. They cannot stand very high temperatures, and they thrive in environments with neutral to slightly acidic pH levels.
Function of a Saprophyte
An example of a substance that is only broken down by saprophytes is lignin, which is a major component in many plants and is what gives trees their tough characteristics. Additionally, a saprophyte is helpful to the ecosystem because as it decomposes the bodies of dead organisms, it recycles and releases nutrients into the environment, making them available for other organisms to use. This is especially important for plant growth.
- Detritivore – An animal that lives off dead and decaying matter.
- Parasite – An organism that lives on another living organism and causes it harm.
- Photosynthesis – The process where green plants and other organisms use sunlight to synthesize their own food from carbon dioxide and water.
1. How do saprophytic fungi acquire nutrients?
A. By digesting food within their stomachs
B. Through photosynthesis
C. By extracellular digestion, performed by hyphae
D. By eating other living organisms
Answer to Question #1 C is correct. Hyphae grow through the dead material and secrete digestive juices in order to break down organic matter and absorb simple products.
2. Which of the following is true of most fungi?
A. They are autotrophic
B. They are multicellular saprophytes
C. They are unicellular saprophytes
D. They are unicellular parasites
Answer to Question #2 B is correct. Most fungi are multicellular, not unicellular, and are saprophytes that decompose matter.
3. Which of the following describes a mycelium?
A. It is a network of connected hyphae
B. It is the digestive juice produced by hyphae
C. It is the digestive tract of a saprophyte
D. It is a parasitic fungus
Answer to Question #3 A is correct. We refer to the mass of hyphae as a mycelium.
The main difference between parasite and saprophyte is that parasite lives on another organism whereas saprophyte is an organism that uses decomposing matter as a food source.
Parasite vs. Saprophyte
The parasite is an organism which depends on other organisms for its nourishment and growth, and that organism is called the host. Saprophyte is an organism which depends on the dead or decaying matter for its food and growth. Parasites use intracellular digestion to get energy. Intracellular metabolism involves both phagocytosis and autophagy. Saprophytes use an extracellular type of digestion. In this process, digestive substances are secreted on the surrounding to break down the organic matter into pure substances, and then degraded substances are absorbed back. Parasites show different relations with their hosts. Sometimes they cause harm to the hosts and even prove lethal for hosts, and sometimes it takes nutrition from the host and gives benefit in return; this type of relation is called a symbiotic relationship. Saprophytes depend only on the dead matter, or decomposing matter so causes no harm. Saprophytes are very beneficial to the ecosystems in the carbon cycle, nitrogen cycle, hydrogen and minerals cycles.
|The parasite is an organism that lives on or into another organism (host) temporarily or permanently and also using it as a source of food.||Saprophyte is an organism which feeds on the decomposing matter of dead organisms.|
|Eukaryotic organism||Both prokaryotic and eukaryotic organism|
|Specialized to parasite on specific hosts||Not strictly specialized|
|Alive host||Decomposing matter of the dead organism|
|Harmful for the hosts, sometimes cause death||Not dangerous for living organisms, beneficial for the environment|
|Type of Digestion|
|Intracellular digestion||Extracellular digestion|
|Wasps, Plasmodium, Calcutta, etc||Bacteria, certain fungi, plants and animals|
What is Parasite?
Parasites vary widely in size and types. Almost 70% of the parasites cannot be seen with naked eyes, for example, malarial parasite, but some worm parasites can reach up to 30 meters in length. The parasite is not a disease itself, but it can spread diseases in hosts. Different parasites have different effects. Unlike predators, parasites do not kill the hosts directly or do not kill it at all. But there is a form of parasitism in which the parasite directly kills its host. This type is called parasitoids. Its example is some species of wasps parasitizing on spiders. This type of relationship is a transient between parasitism and predation. Depending on the relationship between the parasite and its host concerning time and space, parasitism can be of different types. Obligate parasites live at least in one stage while facultative parasites are the free type of organisms, but they find suitable hosts, they switch to the parasitic life. Ectoparasite lives on the surface of its host; skin, feathers, fur or grills whereas endoparasites live inside the host’s body; tissues, cells or body cavity. Some parasites are the ectoparasites at a particular stage of life and then become endoparasites. Temporary parasites spend their lives outside the host but become attached to the host when they need to feed while permanent parasites spend their entire lives within the host’s body. Most of the parasites are specialized to parasites at specific hosts. The obligatory parasites are more specialized than the facultative parasites. In hyperparasitism, parasites are also hosts.
What is Saprophyte?
Saprophytes generally refer to the plants. It can also point to a specific type of orchids and a family of flowering plants called monotropic. Monotropes do not use photosynthesis to make nutrition. Instead, they extract nutrients from dead organic matter. Saprophytic organisms play a significant role in the ecosystem and the circulation of substances in the biosphere. Saprophytes process organic materials from both heterotrophic and heterotrophic microorganisms. This is because of the saprophytic relationship that the ground is not covered with dead organic matter. Some groups of saprophytes decompose complex organic substances to simple substances. For example, proteins are broken down into pure amino acids by breaking the peptide bonds; lipids are broken down into glycerol, and fatty acids by lipase and starch are broken down into simple disaccharides by amylases. Some groups of saprophytes process simple organic substances to inorganic substances. All varieties of the saprophytic organism convert the organic substances formed by autotrophic and consume by the heterotrophic organism into inorganic ones. For any saprophytic nutrition, there are some optimum conditions that must be available; the presence of water, oxygen, neutral or acidic pH and low-medium temperature 1 to 30°C. 80 to 90% of the Fungi are made up of water mass, so they require water content available for nutrition. Similarly, very few saprophytes can survive in anaerobic conditions.
- Parasites are eukaryotic organism whereas saprophytes can be both eukaryotic and prokaryotic organisms.
- Parasites are specialized to parasite on specific organism or hosts whereas saprophytes are not strictly specialized on specific hosts, they can feed on a variety of matter.
- Parasites get their food from the hosts when they are alive whereas saprophytes get their food from the organisms when they are dead.
- Parasites show intracellular digestion whereas saprophytes show extracellular digestion.
- Some parasites develop haustoria to absorb nutrients from the host whereas saprophytes secrete enzymes and degrade organic matter for absorption.
- Parasites prove very harmful and dangerous for the hosts whereas saprophytes do not show any harm for living organisms.
From the above discussion, it has concluded that parasites are most dangerous for other organisms while saprophytes are beneficial for the ecosystem.
What Is A Saprophyte And What Do Saprophytes Feed On
When people think about fungi, they usually think about unpleasant organisms such as poisonous toadstools or those that cause moldy food. Fungi, along with some types of bacteria, belong to a group of organisms called saprophytes. These organisms play an important role in their ecosystem, making it possible for plants to thrive. Find out more about saprophytes in this article.
What is a Saprophyte?
Saprophytes are organisms that can’t make their own food. In order to survive, they feed on dead and decaying matter. Fungi and a few species of bacteria are saprophytes. Examples saprophyte plants include:
- Indian pipe
- Corallorhiza orchids
- Mushrooms and molds
- Mycorrhizal fungi
saprophyte organisms feed, they break down decaying debris left by dead plants and animals. After the debris is broken down, what remains are rich minerals that become part of the soil. These minerals are essential for healthy plants.
What Do Saprophytes Feed On?
When a tree falls in the forest, there may not be anyone there to hear it, but you can be sure that there are saprophytes there to feed on the dead wood. Saprophytes feed on all types of dead matter in all sorts of environments, and their food includes both plant and animal debris. Saprophytes are the organisms responsible for turning food waste you throw into your compost bin into rich food for plants.
You may hear some people refer to exotic plants that live off of other plants, such as orchids and bromeliads, as saprophytes. This isn’t strictly true. These plants often consume live host plants, so they should be called parasites rather than saprophytes.
Additional Saprophyte Information
Here are some features that can help you determine whether an organism is a saprophyte. All saprophytes have these characteristics in common:
- They produce filaments.
- They have no leaves, stems or roots.
- They produce spores.
- They can’t perform photosynthesis.
RESULTS AND DISCUSSION
There was consistently a higher abundance of bacterial than archaeal 16S rRNA gene copy numbers across the pH gradient, as shown by a ratio between archaeal and bacterial copy numbers that ranged between 0.002 and 0.07. We identified two distinct and opposite effects of pH on the abundance of archaeal 16S rRNA gene copies across the pH interval covered by the gradient. Between pH 4.0 and 4.7 (referred to here as the “low-pH” range), the abundance of archaeal 16S rRNA gene copies was on average 8.7 × 103 ± 0.92 × 103 copies g−1 of soil (mean ± standard error ) (Fig. 1). Above this pH range, there was an almost 4-fold decrease in archaeal abundance, with a minimum of only 2.5 × 103 to 2.7 × 103 copies g−1 of soil at pH 5.1 to 5.2. The archaeal abundance then sharply increased with pH between pH 5.1 and 8.3 (referred to here as the “high-pH” range) (P < 0.001), from a minimum of 2.5 × 103 copies g−1 to a maximum of 3.7 × 105 at pH 8 (Fig. 1), resulting in an almost 150-fold increase in archaeal copy numbers. Our further analyses were carried out in the light of this finding by analyzing the archaeal response at pH 4.0 to 4.7 separately from that of archaea found at pH 5.1 to 8.3.
The relationship between soil pH and archaeal abundance. Open circles represent samples with pH 5.1 to 8.3 and filled squares samples with pH 4.0 to 4.7. Only samples in the pH range 5.1 to 8.3 were included in the regression analysis.
The two disparate responses of the archaeal community at low and high pHs are consistent with the recent proposal that some ammonium-oxidizing archaea (AOA) are specialized to the conditions of high-pH environments and others to low pH (20, 33). Even so, the strong positive relationship between soil pH and archaeal abundance at pH 5.1 to 8.3 was surprising. Soil pH is considered one of the major factors regulating the abundance of AOA, and their abundance relative to bacterial ammonium oxidizers is generally higher at low pH (16, 33). Our findings seem to contrast with this. In the low-pH range (pH 4.0 to 4.7), the ratio between archaeal and bacterial copy numbers was on average 0.009 ± 0.0006 (mean ± SE), which decreased to a minimum of 0.002 at pH 5.2. However, above this pH the ratio steadily increased by about 40-fold with higher pH (P < 0.001), reaching a maximum of more than 0.07 at pH 8 (Fig. 2), consistent with other reports of decreasing AOA abundance with decreasing pH (12, 20, 22). A possible explanation for the contradictory results in different reports may be harbored in our results obtained here: the archaeal response to pH is possibly dependent on, and variable with, the span and resolution of the pH range that is examined. We present here one of the most highly resolved and comprehensive ranges of soil pHs so far used within the same study to assess archaeal abundances, and our findings suggest that archaeal abundance responded to pH in two disparate ways along the gradient, probably as a result of some archaea being specialized to the conditions of high-pH environments and others to low pH (20, 33).
The relationship between soil pH and the ratio between archaeal and bacterial copy numbers . The lowest ratio between logarithmically transformed copy numbers (0.55 at pH 5.2) corresponds to a ratio of archeal/bacterial copy numbers of 0.002 and the highest value for the ratio between logarithmically transformed counts (0.85 at pH 8.0) to a ratio of archeal/bacterial copy numbers of 0.07. Open circles represent samples with pH 5.1 to 8.3 and filled squares samples with pH 4.0 to 4.7. Only samples in the pH range of 5.1 to 8.3 were included in the regression analysis. The data on bacterial 16S rRNA gene copy numbers has been previously reported in reference 38.
Another possible reason for the nonuniform response of archaea to soil pH along the gradient is that we quantified the total archaeal community, not constraining our analysis to only the AOA subgroup. A previous study (28) observed a massive decline of the abundance of group 1.1c Crenarchaeota with increased pH (between pH 4.5 and 6.0). Crenarchaeota group 1.1c is also commonly the dominant group of archaea in acidic forest soils (11, 34, 45). It is possible that our observation of decreases in archaeal abundance from pH 4.0 to 4.7 to pH 5.2 (Fig. 1) reflects a reduced abundance of this particular group of archaea, and to our knowledge there is no evidence that these archaea are capable of ammonia oxidation. On the other hand, Crenarchaeota group 1.1c archaea were recently found to have the ability to grow on methanol and methane (9), and observations of archaeal growth in the presence of acetylene (an inhibitor of ammonia oxidation) provides further evidence that archaea in upland soils might not be sustained by NH4+ oxidation alone (22).
The highest pH values in our study correspond well to the pH of seawater (∼pH 8), where archaea are abundant (42) and are at least partly sustained by heterotrophic growth on organic acids (6, 35, 43). In agreement with these findings, the strong positive relationship between the abundance of archaea and pH in the high-pH range (pH 5.1 to 8.3) in this study (Fig. 1) coincided with a slightly reduced leucine uptake per bacterial cell, as suggested by a weak but nonsignificant (P = 0.09) negative relationship between archaeal abundance and specific bacterial growth (ratio between leucine incorporation and bacterial 16S rRNA gene copy numbers). The same relationship became highly significant (P = 0.013) when only samples with pH between 6.0 and 8.3 were included in the analysis (Fig. 3), i.e., the pH range where the increase in the abundance of archaeal 16S rRNA gene copies was most notable (Fig. 1). The same effect was not observed to the same extent for bacterial thymidine incorporation, as the ratio between bacterial leucine uptake and thymidine uptake decreased slightly when archaea became more abundant (P = 0.002; Fig. 4). Thymidine uptake is generally considered to be specific to bacteria (3, 31), while the capacity for uptake of exogenous leucine is a more ubiquitous trait among microorganisms (7, 36). Thus, it cannot be ruled out that archaeal leucine utilization contributed to the observed decrease in bacterial leucine incorporation versus thymidine incorporation (Fig. 4). However, as our study was not designed to elucidate the potential for archaeal amino acid uptake or heterotrophy, we have no direct evidence for this conjecture. Nevertheless, the observations warrant further investigation of the possibility that archaea contribute to the rapid cycling of organic acids not only in marine environments (6) but also in soils of neutral to alkaline pHs. Reinforcing this argument is evidence that the recently isolated archaeal ammonium oxidizer Nitrososphaera viennensis grows more vigorously when the growth medium is enriched with the organic acid pyruvate (46). On a separate note, the potential archaeal uptake of leucine suggests a path forward to estimate archaeal growth rates in soil (21).
The relationship between archaeal abundance and “specific” bacterial leucine incorporation (attomoles leucine incorporated per bacterial 16S rRNA gene copy). There was a weak but nonsignificant (P = 0.09) negative relationship at pH 5.1 to 8.3 which became highly significant (P = 0.013) when only samples with pH between 6.0 and 8.3 were included in the analysis. Open circles represent samples with pH 6.0 to 8.3 and filled squares samples with pH 4.0 to 5.6. The bacterial qPCR data and the bacterial growth rate here used to evaluate the archaeal results were previously reported in reference 38 and reference 39, respectively.
The relationship between archaeal abundance and the ratio of bacterial leucine (Leu) incorporation to thymidine (TdR) incorporation. Open circles represent samples with pH 5.1 to 8.3 and filled squares samples with pH 4.0 to 4.7. Only samples in the pH range of 5.1 to 8.3 were included in the regression analysis. The bacterial growth rates here used to evaluate the archaeal results have been previously reported in a different context in reference 39.
If there is overlap in the use of resources by archaea and bacteria, we might expect to find low archaeal abundance where conditions promote high bacterial growth rates (24, 47). This was not the case. On the contrary, archaea became more abundant at high pH (Fig. 1), where bacterial growth rates were also the highest (39). However, these findings alone do not allow us to exclude the possibility that there are competitive interactions between archaea and bacteria, as the sharp increase in archaeal abundance in the high-pH range might have been even more pronounced in the absence of bacteria.
In contrast, there was a strong negative relationship between fungal growth and archaeal abundance (Fig. 5). That is, when conditions favored high fungal growth rates, archaeal abundance decreased. However, the response was obvious only in the high-pH range (pH 5.1 to 8.3; P < 0.001) whereas there was no correlation between fungal growth and archaeal abundance below pH 5.1. The results contradict observations suggesting that mycorrhizal fungi might actually promote growth of archaea (8, 9, 11). However, those observations are derived from experiments using boreal pine forest soils with pH matching the low-pH range in our study (pH 4.0 to 4.7), i.e., the range where archaeal abundance was not negatively affected by fungal growth (Fig. 5). In contrast, another study (24) found a pH-independent decrease in archaeal abundance, as well as in the ratio between archaea and bacteria, in direct proximity to mycorrhizal roots and mycorrhizal hyphae in a forest soil with pH 5.5. Our observation of decreased archaeal abundance at high fungal growth rates is consistent with those findings.
The relationship between fungal growth and archaeal abundance. Open circles represent samples with pH 5.1 to 8.3 and filled squares samples with pH 4.0 to 4.7. Only samples in the pH range of 5.1 to 8.3 were included in the regression analysis. The fungal growth rates here used to evaluate the archaeal results have been previously reported in a different context (39).
The ratio between archaeal and bacterial 16S rRNA gene copy numbers also decreased considerably with increasing fungal growth rates in the high-pH range (P < 0.001, Fig. 6), suggesting that archaea rather than bacteria are the prokaryotes most strongly affected by interactions with fungi. However, the results should be interpreted with caution, as autocorrelation between the measured biological and environmental factors makes the observed correlative relationships between microbial groups difficult to assign to causal factors. Moreover, that competitive interaction between fungi and bacteria appears to explain their pH dependencies (40) complicates the picture further. It should also be noted that the acetate incorporation assay used to estimate fungal growth rates measures only the growth of saprotrophic fungi and not that of mycorrhizal fungi. It can be speculated that mycorrhizal fungi provide growth factors or an environment that favors archaeal growth, while the same is not true for saprotrophic fungi. Furthermore, the correlative measures of archaeal abundance and fungal and bacterial growth should be interpreted with some caution, since we do not have any information on archaeal growth rates in the same samples. However, lacking a better measure of the archaeal presence, this is a valuable initial assessment.
The relationship between fungal growth and the ratio between archaeal and bacterial abundance (log10 16S rRNA gene copy numbers g−1 soil). Open circles represent samples with pH 5.1 to 8.3 and filled squares samples with pH 4.0 to 4.7. Only samples in the pH range of 5.1 to 8.3 were included in the regression analysis. The bacterial qPCR data and the fungal growth rates here used to evaluate the archaeal results were previously reported in reference 38 and reference 39, respectively.
In conclusion, our results show that soil pH is an important factor that regulates archaeal abundance in the terrestrial environment. Interestingly, however, the archaeal response to pH was not uniform and our results implicitly suggest that two disparate influences of pH on the archaeal community were present. This might possibly be a reflection of variations in the archaeal community composition along the gradient, with some archaea being adapted to acidic conditions and others to neutral to slightly alkaline conditions. This conclusion is reinforced by observations of contrasting outcomes of the (competitive) interactions between archaea, bacteria, and fungi toward the lower and higher ends of the examined pH gradient. Our assessment of the influence of soil pH on the distribution of archaeal abundance was devised to factor in the whole archaeal domain, and our results contrast with what is growing to be a consensus regarding the relationship between AOA and soil pH. This, combined with recent finding of alternative archaeal nutritional strategies, emphasizes a need to reconsider the ecology and resource use of soil archaea—they may be more diverse than is currently recognized.