ps 3 – reaction centers

Two photosynthetic reaction centers are arranged in tandem in photosynthesis of algae and plants

In green algae about eight photons are required (quantum requirement: pho- tons absorbed per molecule O₂ produced) for the photosynthetic water split- ting (section 2.4). Instead of the term quantum requirement, one often uses the reciprocal term quantum yield (molecules of O₂ produced per photon absorbed). According to the color of irradiated light (action spectrum) the quantum yield dropped very sharply when algae were illuminated with red light above a wavelength of 680 nm (Fig. 3.14). This effect, named “red drop,” remained unexplained since algae contain chlorophyll, which absorbs light at 700 nm. Robert Emerson and coworkers (USA) solved this problem in 1957 when they observed that the quantum yield in the spectral range above 680 nm increased dramatically when algae were illuminated with orange light (650 nm) and red light simultaneously. Then the quantum yield was higher than the sum of both yields when irradiated separately with the light of each wavelength. This Emerson effect led to the conclusion that two differ- ent reaction centers are involved in photosynthesis of green algae (and also of cyanobacteria and higher plants). In 1960 Robert Hill (Cambridge, UK) postulated a reaction scheme (Fig. 3.15) in which two reaction centers are

Figure 3.17 Schematic presentation of the localization of the photosynthetic complexes and the H^-ATP synthase in the thylakoid membrane. Transport of electrons between PS II and the cytochrome-b₆/f complex is mediated by plastohydroquinone (PQH₂), and that between the cytochrome-b₆/f complex and PS I by plastocyanin (PC). Water splitting occurs on the luminal side of the membrane, and the formation of NADPH and ATP on the stromal side. The electrochemical gradient of protons pumped into the lumen drives ATP synthesis. The number of protons transported to the lumen during electron transport and the proton requirement of ATP synthesis is not known (section 4.4).

Arranged in tandem and connected by an electron transport chain containing cytochrome-b₆ and cytochrome-f (cytochrome-f is a cytochrome of the c type; see section 3.7). Light energy of 700 nm was sufficient for the excitation of reaction center I, whereas excitation of the other reaction center II required light of higher energy with a wavelength of 680 nm. The electron flow accord- ing to the redox potentials of the intermediates shows a zigzag, leading to the name Z scheme. The numbering of the two photosystems corresponds to the sequence of their discovery. Photosystem II (PS II) can use light up to a wave- length of 680 nm, whereas photosystem I (PS I) can utilize light with a wave- length up to 700 nm. The sequence of the two photosystems makes it possible that at PS II a very strong oxidant is generated for the oxidation of water and at PS I a very strong reductant is produced for the reduction of NADP^ (see also Fig. 3.3).

Figure 3.16 gives an overview of electron transport through the photo- synthetic complexes; the carriers of electron transport are drawn according to their electric potential (see also Fig. 3.11). Figure 3.17 shows how the photosynthetic complexes are arranged in the thylakoid membrane. There is a potential difference of about 1.2 volt between the process of water oxi- dation and NADP^ reduction. The absorbed photons of 680 and 700 nm together correspond to a total potential difference of 3.45 volt (see section 2.2, equation 2.7). Thus, only about one-third of the energy of the pho- tons absorbed by the two photosystems is used to transfer electrons from

water to NADP^. In addition to this, about one-eighth of the light energy absorbed by the two photosystems is conserved by pumping protons into the lumen of the thylakoids via PS II and the cytochrome-b₆/f complex (Fig. 3.17). This proton transport leads to the formation of a proton gradi- ent between the lumen and the stroma sp ace. An H^-ATP synthase, also located in the thylakoid membrane, uses the energy of the proton gradient to synthesize ATP. Thus about half the absorbed light energy of the two photosystems is not used for chemical work but is dissipated as heat. The significance of the loss of energy as heat during photosynthetic electron transport has been discussed in section 2.3.

Water is split by photosystem II

The groups of Horst Witt and Wolfgang Saenger (both in Berlin) resolved the three-dimensional structure of PS II by X-ray structure analysis of crys- tals from the PS II of the thermophilic cyanobacteria Thermosynechococcus elongatis. The subsequent X-ray structure analysis of PS I revealed that PS II and PS I are constructed after the same basic principles as the reaction

centers of purple bacteria (section 3.4). This, and sequence analyses, clearly demonstrate that all these photosystems have a common origin. Thus PS II also has a chl-a pair in the center, although the distance between the two molecules is so large that probably only one of the two chl-a molecules reacts with the exciton. Two arms, each with one chl-a and one pheophy- tin molecule, are connected with this central pair as in the purple bacteria shown in Figure 3.10. Also in the cyanobacteria, only one of these arms appears to be involved in the electron transport.

In contrast to the bacterial reaction center the excitation of the reaction center results in an electron transfer via the chl-a monomer to pheophytin (Phe), and from there to a tightly bound plastoquinone (Q_A), thus forming a semiquinone radical (Fig. 3.18). The electron is then further transferred to a loosely bound plastoquinone (Q_B). This plastoquinone (PQ) (Fig. 3.19) accepts two electrons and two protons one after the other and is thus reduced to hydroquinone (PQH₂). The hydroquinone is released from the photosynthesis complex and may be regarded as the final product of photosystem II. This sequence, consisting of a transfer of a single electron between (chl-a)₂ and Q_A and the transfer of two electrons between Q_A and Q_B, corresponds to the reaction sequence shown for Rb. sphaeroides (Fig. 3.11). The only difference is that the quinones are ubiquinone or menaqui- none in bacteria and plastoquinone in photosystem II.

However, the similarity between the reaction sequence in PS II and the photosystem of the purple bacteria applies only to the electron acceptor region. The electron donor function in PS II of plants is completely differ- ent from that in purple bacteria. The electron deficit in (chl-a)₂ ^ caused by non-cyclic electron transport is compensated for by electrons derived from the oxidation of water. In the transport of electrons from water to chloro- phyll manganese cations and a tyrosine residue are involved. The (chl-a)₂ ^ radical with a redox potential of about 1.1 volt is such a strong oxidant that it can withdraw an electron from a tyrosine residue in the protein of the reaction center and a tyrosine radical remains. This reactive tyrosine residue is often designated as Z. The electron deficit in the tyrosine radical is restored by oxidation of a manganese ion (Fig. 3.20). The PS II com- plex contains several manganese ions, probably four, which are close to each other. This arrangement of Mn ions is called the Mn cluster. The Mn cluster depicts a redox system that can take up and release four electrons. During this process the Mn ions probably change between the oxidation state Mn^3 and Mn^4. To liberate one molecule of O₂ from water, the reaction center must with- draw four electrons and thus capture four excitons. The time differences between the capture of the single exciton in the reaction center depends on the intensity of illumination. If oxidation of water were to proceed stepwise, oxygen radicals could be formed as intermediary products, especially at low light intensities. Oxygen radicals have a destructive effect on biomolecules such as lipids and proteins (section 3.10). The water splitting machinery of the Mn clusters minimizes the formation of oxygen radical intermediates by supplying the reaction center via tyrosine with four electrons one after the other (Fig. 3.20). The Mn cluster is transformed during this transfer from the ground oxidation state stepwise to four different oxidation states (these have been designated as S₀ and S₁–S₄).

Experiments by Pierre Joliot (France) and Bessel Kok (USA) presented evidence that the water splitting apparatus can be in five different oxida- tion states (Fig. 3.21). When chloroplasts that were kept in the dark were then illuminated by a series of light pulses, an oscillation of the oxygen release was observed. Whereas after the first two light pulses almost no O₂ was released, the O₂ release was maximal after three pulses and then after a further four pulses, and so on. An increasing number of light pulses, how- ever, dampened the oscillation. This can be explained by pulses that do not cause excitation of PS II and thus desynchronize the oscillation. In dark- ened chloroplasts the water splitting apparatus is in the S₁ state. After the fourth oxidation state (S₄) has been reached, O₂ is released in one reaction and the Mn cluster returns to its ground oxidation state (S₀). During this reaction, protons from water are released to the lumen of the thylakoids. The formal description of this reaction is:

Figuratively speaking, the four electrons needed in the reaction center are loaned in advance by the Mn cluster and then repaid at one stroke by oxidizing water to synthesize one oxygen molecule. In this way the Mn clus- ter minimizes the formation of oxygen radicals in photosystem II. Despite this safety device, still some oxygen radicals are formed in the PS II com- plex which damage the proteins of the complex. The consequences will be discussed in section 3.10.

Photosystem II complex is very similar to the reaction center in purple bacteria

Photosystem II is a complex consisting of at least 20 different subunits (Table 3.2), only two of which are involved in the actual reaction center. For the sake of simplicity the scheme of the PS II complex shown in Fig.

3.22 contains only some of these subunits. The PS II complex is surrounded by an antenna consisting of light harvesting complexes (Fig. 2.13).

The center of the PS II complex is a heterodimer consisting of the sub- units D₁ and D₂ with six chl-a, two pheophytin, two plastoquinone, and one to two carotenoid molecules bound to it. The D₁ and D₂ proteins are homologous to each other and also to the L proteins and M proteins from the reaction center of the purple bacteria (section 3.4). As in purple bacteria, only the pheophytin molecule bound to the D₁ protein of PS II is involved in electron transport. Q_A is bound to the D₂ protein, whereas Q_B is bound to the D₁ protein. The Mn cluster is probably enclosed by both the D₁ and D₂ proteins. The tyrosine that is reactive in electron transfer is a constituent

of D₁. The subunits O, P, Q stabilize the Mn cluster. The two subunits CP 43 and CP 47 (CP means chlorophyll protein) each bind about 15 chloro- phyll molecules and form the core complex of the antenna shown in Figure

2.10. CP 43 and CP 47 flank both sides of the D₁-D₂ complex. Cyt-b₅₅₉ does not seem to be involved in the electron transport of PS II; possibly its func- tion is to protect the PS II complex from light damage. The inner and outer light harvesting complexes of LHC II are arranged at the periphery.

The D₁ protein of the PS II complex has a high turnover; it is constantly being resynthesized. It seems that the D₁ protein wears out during its func- tion, perhaps through damage by oxygen radicals, which still occurs despite all the protection mechanisms. It has been estimated that the D₁ protein is replaced after 10⁶ to 10⁷ catalytic cycles of the PS II reaction center.

A number of compounds that are similar in their structure to plasto- quinone can block the plastoquinone binding site at the D₁ protein, caus- ing inhibition of photosynthesis. Such compounds are used as weed killers (herbicides). Before the effect of these compounds is discussed in detail, some general aspects of the application of herbicides shall be introduced.

Table 3.4: Protein components of photosystem I (list not complete)

Mechanized agriculture usually necessitates the use of herbicides

About 50% of the money spent worldwide for plant protection is expended for herbicides. The high cost of labor is one of the main reasons for using herbicides in agriculture. It is cheaper and faster to keep a field free of weeds by using herbicides rather than manual labor. Weed control in agriculture is necessary not only to decrease harvest losses by weed competition, but also because weeds hinder the operation of harvesting machineries; fields free of weeds are a prerequisite for a mechanized agriculture. The herbicides usu- ally block a specific reaction of the plant metabolism and have a low toxic- ity for animals and humans. A large number of herbicides (examples will be given at the end of this section) inhibit photosystem II by being antagonists to plastoquinone. To achieve substantial inhibition the herbicide molecule has to bind to most of the many photosynthetic reaction centers. To be effective, 125 to 4,000 g of these herbicides have to be applied per hectare.

In an attempt to reduce the amount of herbicides applied to the soil, new efficient herbicides have been developed that inhibit key biosynthetic processes such as the synthesis of fatty acids, amino acids (sections 10.1 and 10.4), carotenoids, or chlorophyll. There are also herbicides that act as analogues of phytohormones or mitosis inhibitors. Some of these herbi- cides are effective with amounts as low as 5 g per hectare.

Some herbicides are taken up only by the roots and others by the leaves. To keep the railway tracks free of weeds, nonselective herbicides are employed, which destroy the complete vegetation. Nonselective herbicides are also used in agriculture, e.g., to combat weeds in citrus plantations. In the latter case, herbicides are applied that are only taken up by the leaves to combat herbaceous plants at the ground level. Especially interesting are selec- tive herbicides that combat only weeds and effect cultivars as little as possible (sections 12.2 and 15.3). Selectivity can be due to different uptake efficien- cies of the herbicide in different plants, different sensitivities of the metabo- lism towards the herbicide, or different ability of the plants to detoxify the herbicide. Important mechanisms that plants utilize to detoxify herbicides and other foreign compounds (xenobiotics) are the introduction of hydroxyl groups by P-450 monooxygenases (section 18.2) and the formation of glu- tathione conjugates (section 12.2). Selective herbicides have the advantage that weeds can be destroyed at a later growth stage of the cultivars where the dead weeds form a mulch layer conserving water and preventing erosion.

In some cases, the application of herbicides has led to the evolution of herbicide-resistant plant mutants (section 10.4). Conventional breeding has used such mutated plants to generate herbicide-resistant cultivars. In contrast to the occurrence of herbicide resistance by accidental mutations, nowadays genetic engineering is employed on a very large scale to generate cultivars which are resistant to a certain herbicide, allowing weed control in the presence of the growing cultivar (section 22.6)

A large number of herbicides inhibit photosynthesis: the urea deriva- tive DCMU (Diuron, DuPont), the triazine Atrazine (earlier Ciba Geigy), Bentazon (BASF) (Fig. 3.23), and many similar compounds function as herbicides by binding to the plastoquinone binding site on the D₁ pro- tein and thus blocking the photosynthetic electron transport. Nowadays, DCMU is not often used, as the required dosage is high and its degradation is slow. It is, however, often used in the laboratory to inhibit photosynthe- sis in an experiment (e.g., of leaves or isolated chloroplasts). Atrazine acts selectively: maize plants are relatively insensitive to this herbicide since they have a particularly efficient mechanism for its detoxification (section 12.2). Because of its relatively slow degradation in the soil, the use of Atrazine has been restricted in some countries, e.g., Germany. In areas where cer- tain herbicides have been used continuously over the years, some weeds have become resistant to these herbicides. In some cases, the resistance can be traced back to mutations resulting in a single amino acid change in the D₁-proteins. These changes do not markedly affect photosynthesis of these weeds, but they do decrease binding of the herbicides to the D₁-protein.

The cytochrome-b₆/f complex mediates electron transport between photosystem II and photosystem I Iron atoms in cytochromes and in iron-sulfur centers have a central function as redox carriers

Cytochromes occur in all organisms except a few obligate anaerobes. These are proteins to which one to two tetrapyrrole rings are bound. These tetrapyrroles are very similar to the chromophores of chlorophylls. However, chlorophylls contain Mg^ as the central atom in the tetrapyr- role, whereas the cytochromes have an iron atom (Fig. 3.24). The tetrapy- rrole ring of the cytochromes with iron as the central atom is called the heme. The bound iron atom can change between the oxidation states Fe^ and Fe^ so that cytochromes function as a one-electron-carrier, in con- trast to quinones, NAD(P) and FAD, which transfer two electrons together with protons.

Cytochromes are divided into three main groups, the cytochromes-a, –b, and –c. These correspond to heme-a, –b, and –c. Heme-b may be regarded as the basic structure (Fig. 3.24). In heme-c the -SH-group of a cysteine is added to each of the two vinyl groups of heme-b. In this way heme-c is

covalently bound by a sulfur bridge to the protein of the cytochrome. Such a mode of covalent binding has already been shown for phycocyanin in Figure 2.15, and there is actually a structural relationship between the correspond- ing apoproteins. In heme-a (not shown) an isoprenoid side chain consisting of three isoprene units is attached to one of the vinyl groups of heme-b. This side chain functions as a hydrophobic membrane anchor, similar to that found in quinones (Figs. 3.5 and 3.19). Heme-a is mentioned here only for the sake of completeness. It plays no role in photosynthesis, but it does have a function in the mitochondrial electron transport chain (section 5.5).

The iron atom in the heme can form up to six coordinative bonds. Four of these bonds are formed with the nitrogen atoms of the tetrapyrrole ring. This ring has a planar structure. The two remaining bonds of the Fe atom coordinate with two histidine residues, which are positioned vertically to the tetrapyrrole plane (Fig. 3.25). Cyt-f (f  foliar, in leaves) contains, like cyt-c, one heme-c and therefore belongs to the c-type cytochromes. In cyt-f one bond of the Fe atom coordinates with the terminal amino group of the protein and the other with a histidine residue.

Iron-sulfur centers are of general importance as electron carriers in elec- tron transport chains and thus also in photosynthetic electron transport. Cysteine residues of proteins within iron-sulfur centers (Fig. 3.26) are coor- dinatively or covalently bound to Fe atoms. These iron atoms are linked to each other by S-bridges. Upon acidification of the proteins, the sulfur between the Fe atoms is released as H₂S and for this reason it has been called labile sulfur. Iron-sulfur centers occur mainly as 2Fe-2S or 4Fe-4S centers. The Fe atoms in these centers are present in the oxidation states

Fe^ and Fe^. Irrespective of the number of Fe atoms in a center, the oxidized and reduced state of the center differs only by a single charge. For this reason, iron-sulfur centers can take up and transfer only one electron. Various iron-sulfur centers have very different redox potentials, depending on the surrounding protein.

The electron transport by the cytochrome-b₆/f complex is coupled to a proton transport

Plastohydroquinone (PQH₂) formed by PS II diffuses through the lipid phase of the thylakoid membrane and transfers its electrons to the cytochrome-b₆/f complex (Fig. 3.17). This complex then transfers the electrons to plastocy- anin, which is thus reduced. Therefore the cytochrome-b₆/f complex has also been called plastohydroquinone-plastocyanin oxidoreductase. Plastocyanin is a protein with a molecular mass of 10.5 kDa, containing a copper atom, which is coordinatively bound to one cysteine, one methionine, and two histidine residues of the protein (Fig. 3.27). This copper atom alternates between the oxidation states Cu^ and Cu^ and thus is able to take up and transfer one electron. Plastocyanin is soluble in water and is located in the thylakoid lumen.

Electron transport through the cyt-b₆/f complex proceeds along a poten- tial difference gradient of about 0.4 V (Fig. 3.16). The energy liberated by the transfer of the electron down this redox gradient is conserved by trans- porting protons to the thylakoid lumen. The cyt-b₆/f complex is a mem- brane protein consisting of at least eight subunits. The main components of this complex are four subunits: cyt-b₆, cyt-f, an iron-sulfur protein called Rieske protein after its discoverer, and a subunit IV. Additionally, there are some smaller peptides and a chlorophyll and a carotenoid of unknown function. The Rieske protein has a 2Fe-2S center with the very positive redox potential of 0.3 V, untypical of such iron-sulfur centers.

The cyt-b₆/f complex has an asymmetric structure (Fig. 3.28). Cyt-b₆ and subunit IV span the membrane. Cyt-b₆ containing two heme-b molecules is almost vertically arranged to the membrane and forms a redox chain across

one iron-sulfur protein. The amino acid sequence of cyt-b in the cyt-b/c₁ complex of bacteria and in mitochondria corresponds to the sum of the sequences of cyt-b₆ and the subunit IV in the cyt-b₆/f complex. Apparently during evolution the cyt-b gene was cleaved into two genes, for cyt-b₆ and subunit IV. Whereas in plants the cyt-b₆/f complex reduces plastocyanin, the cyt-b/c₁ complex of bacteria and mitochondria reduces cyt-c. Cyt-c is a very small cytochrome molecule that is water-soluble and, like plasto- cyanin, transfers redox equivalents from the cyt-b₆/f complex to the next complex along the aqueous phase. In cyanobacteria, which also possess a cyt-b₆/f complex, the electrons are transferred from this complex to photo- system I via cyt-c instead of plastocyanin. The great similarity between the cyt-b₆/f complex in plants and the cyt-b/c₁ complexes in bacteria and mito- chondria suggests that these complexes have basically similar functions in photosynthesis and in mitochondrial oxidation: they are proton transloca- tors that are driven by a hydroquinone-plastocyanin (or -cyt-c) reductase.

The interplay of PS II and the cyt-b₆/f complex electron transport causes the transport of protons from the stroma space to the thylakoid lumen. The principle of this transport is explained in the schematic presentations of Figures 3.28 and 3.29. A crucial point is that the reduction and oxida- tion of the quinone occur at different sides of the thylakoid membrane. The required protons for the reduction of PQ (Q_b) by the PS II complex are taken up from the stroma space. Subsequently PQH₂ diffuses across the lipid phase of the membrane to the binding site in the lumenal region of the cyt-b₆/f complex where it is oxidized by the Rieske protein and cyt-f to yield reduced plastocyanin. The protons of this reaction are released into the thylakoid lumen. According to this scheme, the capture of four excitons by the PS II complex transfers four protons from the stroma space to the lumen. In addition four protons produced during water splitting by PS II are released into the lumen as well.

The number of protons pumped through the cyt-b₆/f complex can be doubled by a Q-cycle

Studies with mitochondria indicated that during electron transport through the cyt-b/c₁ complex, the number of protons transferred per transported electron is larger than four (Fig. 3.29). Peter Mitchell (Great Britain), who established the chemiosmotic hypothesis of energy conservation (section 4.1), also postulated a so-called Q-cycle, by which the number of trans- ported protons for each electron transferred through the cyt-b/c₁ complex is doubled. It later became apparent that the Q-cycle also has a role in pho- tosynthetic electron transport.

Figure 3.30 shows the principle of Q-cycle operation in the photosyn- thesis of chloroplasts. The cyt–b₆/f complex contains two different bind- ing sites for conversion of quinones, one located at the stromal side and the other at the luminal side of the thylakoid membrane (Fig. 3.28). The plastohydroquinone (PQH₂) formed in the PS II complex is oxidized by the Rieske iron-sulfur center at the binding site adjacent to the lumen. Due to its very positive redox potential, the Rieske protein tears off one electron from the plastohydroquinone. Because its redox potential is very negative, the remaining semiquinone is unstable and transfers its electron to the first heme-b of the cyt-b₆ (b_p) and from there to the other heme-b (b_n), thus rais- ing the redox potential of heme b_n to about –0.1 V. In this way a total of four protons are transported to the thylakoid lumen per two molecules of plastohydroquinone oxidized. Of the two plastoquinone molecules (PQ) formed, only one molecule returns to the PS II complex.

The other PQ dif- fuses away from the cyt-b₆/f complex through the lipid phase of the mem- brane to the stromal binding site of the cyt-b₆/f complex to be reduced via semiquinone to hydroquinone by the high reduction potential of heme-b_n. This is accompanied by the uptake of two protons from the stromal space. The hydroquinone thus regenerated diffuses through the membrane back to the luminal binding site where it is oxidized in turn by the Rieske protein, and so on. In total, the number of transported protons is doubled by the Q-cycle (1/2  1/4  1/8  1/16  1/n  1). The fully operating Q-cycle transports four electrons through the cyt-b₆/f complex which results in total to the transfer of eight protons from the stroma to the lumen. The func- tion of this Q-cycle in mitochondrial oxidation is now undisputed, while its function in photosynthetic electron transport is still a matter of contro- versy. The analogy of the cyt-b₆/f complex to the cyt-b/c₁ complex suggests that the Q-cycle also plays an important role in chloroplasts. So far, the operation of a Q-cycle in plants has been observed mainly under low light conditions. The Q-cycle is perhaps suppressed by a high proton gradient generated across the thylakoid membrane, for instance, by irradiation with high light intensity. In this way the flow of electrons through the Q-cycle could be adjusted to the energy demand of the plant cell.

Photosystem I reduces NADP^

Plastocyanin that has been reduced by the cyt-b₆/f complex diffuses through the lumen of the thylakoids, binds to a positively charged binding site of PS I, transfers its electron, and the resulting oxidized form diffuses back to the cyt-b₆/f complex (Fig. 3.31).

Also the reaction center of PS I with an absorption maximum of 700 nm contains a chlorophyll pair (chl-a)₂ (Fig. 3.31). As in PS II, the excitation caused by a photon reacts probably with only one of the two chlorophyll molecules. The resulting (chl-a)₂ ^ is then reduced by plastocyanin. It is assumed that (chl-a)₂ transfers its electron to a chl-a monomer (A₀), which then transfers the electron to a strongly bound phylloquinone (Q) (Fig. 3.32). Phylloquinone contains the same phytol side chain as chl-a and its function corresponds to Q_A in PS II. The electron is transferred from the semiphylloquinone to an iron-sulfur center named F_X. F_X is a 4Fe-4S center

with a very negative redox potential. It transfers one electron to two other 4Fe-4S centers (F_A, F_B), which in turn reduce ferredoxin, a protein with a molecular mass of 11 kDa with a 2Fe-2S center. Ferredoxin also takes up and transfers only one electron. The reduction occurs at the stromal side of the thylakoid membrane. For this purpose, the ferredoxin binds at a posi- tively charged binding site on subunit D of PS I (Fig. 3.33). The reduction of NADP^ by ferredoxin, catalyzed by ferredoxin-NADP reductase, yields NADPH as an end product of the photosynthetic electron transport.

The PS I complex consists of at least 17 different subunits, of which some are shown in Table 3.4. The center of the PS I complex is a heterodimer (as is the center of PS II) consisting of subunits A and B (Fig. 3.33). The molec- ular masses of A and B (each 82–83 kDa) correspond approximately to the sum of the molecular masses of the PS II subunits D₁ and CP₄₃, and D₂ and CP₄₇, respectively (Table 3.2). In fact, both subunits A and B have a dou- ble function. Like D₁ and D₂ in PS II, they bind chromophores (chl-a) and redox carriers (phylloquinone, FeX) of the reaction center and, additionally, they contain about 100 chl-a molecules as antennae pigments. Thus, the het- erodimer of A and B represents the reaction center and the core antenna as well. The three-dimensional structure of photosystem I in cyanobacteria, green algae and plants has been resolved. The principal structure of pho- tosystem I, with a central pair of chl-a molecules and two branches, each with two chlorophyll molecules, is very similar to photosystem II and to the bacterial photosystem (Fig. 3.10). It has not been definitely clarified whether both or just one of these branches are involved in the electron transport. The Fe-S-centers F_A and F_B are ascribed to subunit C, and subunit F is con- sidered to be the binding site for plastocyanin.

The light energy driving the cyclic electron transport of PS I is only utilized for the synthesis of ATP

Besides the noncyclic electron transport discussed so far, cyclic electron transfer can also take place in which the electrons from the excited pho- tosystem I are transferred back to the ground state of PS I, probably via the cyt-b₆/f complex (Fig. 3.34). The energy thus released is used only for the synthesis of ATP, and NADPH is not formed. This electron transport is termed cyclic photophosphorylation. In intact leaves, and even in iso- lated intact chloroplasts, it is quite difficult to differentiate experimentally between cyclic and non-cyclic photophosphorylation. It has been a matter of debate as to whether and to what extent cyclic photophosphorylation occurs in a leaf under normal physiological conditions. Recent evaluations of the proton stoichiometry of photophosphorylation (see section 4.4) sug- gest that the yield of ATP in noncyclic electron transport is not sufficient for the requirements of CO₂ assimilation, and therefore cyclic photophos- phorylation seems to be required to synthesize the lacking ATP. Moreover, cyclic photophosphorylation must operate at very high rates in the bundle sheath chloroplasts of certain C₄ plants (section 8.4). These cells have a high demand for ATP and they contain high PS I activity but very little PSPresumably, the cyclic electron flow is governed by the redox state of theacceptor of the photosystem in such a way that by increasing the reduction of the NADP system, and consequently of ferredoxin, the diversion of the

electrons in the cycle is enhanced. The function of cyclic electron transport is probably to adjust the rates of ATP and NADPH formation according to the plant’s demand.

Despite intensive investigations, the pathway of electron flow from PS I to the cyt-b₆/f complex in cyclic electron transport remains unresolved. It has been proposed that cyclic electron transport is structurally separated from the linear electron transport chain in a super complex. Most experi- ments on cyclic electron transport have been carried out with isolated thy- lakoid membranes that catalyze only cyclic electron transport when redox mediators, such as ferredoxin or flavin adenine mononucleotide (FMN, Fig. 5.16), have been added. Cyclic electron transport is inhibited by the antibiotic antimycin A. It is not clear at which site this inhibitor functions. Antimycin A does not inhibit noncyclic electron transport.

Surprisingly, proteins of the NADP dehydrogenase complex of the mitochondrial respiratory chain (section 5.5) have also been identified in the thylakoid membrane of chloroplasts. The function of these proteins in chloroplasts is still not known. The proteins of this complex occur very fre- quently in chloroplasts from bundle sheath cells of C₄ plants, which have little PS II but a particularly high cyclic photophosphorylation activity (section 8.4). These observations raise the possibility that in cyclic electron transport the flow of electrons from NADPH or ferredoxin to plastoqui- none proceeds via a complex similar to the mitochondrial NADH dehydro- genase complex. As will be shown in section 5.5, the mitochondrial NADH dehydrogenase complex transfers electrons from NADH to ubiquinone. Results indicate that an additional pathway for a cyclic electron transport exists in which electrons are directly transferred via a plastoquinone reduct- ase from ferredoxin to plastoquinone.

In the absence of other acceptors electrons can be transferred from photosystem I to oxygen

When ferredoxin is very highly reduced, it is possible that electrons are

transferred from PS I to oxygen to form superoxide radicals (•O^) (Fig.

3.35). This process is called the Mehler reaction. The superoxide radical reduces metal ions present in the cell such as Fe^3 and Cu^2 (M^n):

The hydroxyl radical (•OH) is a very aggressive substance and damages enzymes and lipids by oxidation. The plant cell has no protective enzymes against •OH. Therefore it is essential that a reduction of the metal ions be prevented by rapid elimination of •O^ by superoxide dismutase. But hydrogen peroxide (H₂O₂) also has a damaging effect on many enzymes. To prevent such damage, hydrogen peroxide is eliminated by an ascorbate

peroxidase located in the thylakoid membrane. Ascorbate, an important anti- oxidant in plant cells (Fig. 3.36), is oxidized by this enzyme and converted to the radical monodehydroascorbate, which is spontaneously reconverted by photosystem I to ascorbate via reduced ferredoxin. Monodehydroascorbate can be also reduced to ascorbate by an NAD(P)H-dependent monodehy- droascorbate reductase that is present in the chloroplast stroma and the cytosol.

As an alternative to the preceding reaction, two molecules of mono- dehydroascorbate can dismutate to ascorbate and dehydroascorbate. Dehydroascorbate is reconverted to ascorbate by reduction with glutath- ione in a reaction catalyzed by dehydroascorbate reductase present in the stroma (Fig. 3.37). Glutathione (GSH) occurs as an antioxidant in all plant

cells (section 12.2). It is a tripeptide composed of the amino acids glutamate, cysteine, and glycine (Fig. 3.38). Oxidation of GSH results in the forma- tion of a disulfide (GSSG) between the cysteine residues of two glutathione molecules. Reduction of GSSG is catalyzed by a glutathione reductase with NADPH as the reductant (Fig. 3.37).

The major function of the Mehler-ascorbate-peroxidase cycle is to dis- sipate excessive excitation energy of photosystem I as heat. The absorption of a total of eight excitons via PS I results in the formation of two super- oxide radicals and two molecules of reduced ferredoxin, the latter serving as a reductant for eliminating H₂O₂ (Fig. 3.35). The transfer of electrons to oxygen by the Mehler reaction is a reversal of the water splitting of PS

II. As will be discussed in the following section, the Mehler reaction occurs when ferredoxin is very highly reduced. The only gain of this reaction is the generation of a proton gradient from electron transport through PS II and the cyt-b₆/f complex. This proton gradient can be used for the synthesis of ATP if ADP is present. But since there is usually a shortage in ADP under the conditions of the Mehler reaction, it mostly results in the formation of a high pH gradient. A feature common to the Mehler reaction and cyclic electron transport is that there is no net production of NADPH. For this reason, electron transport via the Mehler reaction has been termed pseudo- cyclic electron transport.

Yet another group of antioxidants was recently found in plants, the so- called peroxiredoxins. These proteins, comprising -SH groups as redox car- riers, have been known in the animal world for some time. Ten different peroxiredoxin genes have been identified in the model plant Arabidopsis. Peroxiredoxins, being present in chloroplasts as well as in other cell com- partments, differ from the aforementioned antioxidants glutathione and ascorbate in that they reduce a remarkably wide spectrum of peroxides, such as H₂O₂, alkylperoxides, and peroxinitrites. In chloroplasts, oxidized peroxiredoxins are reduced by photosynthetic electron transport of photo- system I with ferredoxin and thioredoxin as intermediates.

Instead of ferredoxin, PS I can also reduce methylviologen. Methylviologen, also called paraquat, is used commercially as a herbicide

(Fig. 3.39). The herbicidal effect is due to the reduction of oxygen to super- oxide radicals. Additionally, paraquat competes with dehydroascorbate for the reducing equivalents provided by photosystem I. Therefore, in the pres- ence of paraquat, ascorbate is no longer regenerated from dehydroascor- bate and the ascorbate peroxidase reaction can no longer proceed. The increased production of superoxide and decreased detoxification of hydro- gen peroxide in the presence of paraquat causes severe oxidative damage to mesophyll cells, noticeable by a bleaching of the leaves. In the past, paraquat has been used to destroy marijuana fields in South America.

Regulatory processes control the distribution of the captured photons between the two photosystems

Linear photosynthetic electron transport through the two photosystems requires the even distribution of the captured excitons between them. As discussed in section 2.4, the excitons are transferred preferentially to the chromophore which requires the least energy for excitation. Photosystem I (P₇₀₀) being on a lower energetic level than PS II (Fig. 3.16) requires less energy for excitation than photosystem II (P₆₈₀). In an unrestricted compe- tition between the two photosystems, excitons would primarily be directed to PS I. Due to this imbalance, the distribution of the excitons between the two photosystems must be regulated. The spatial separation of PS I and PS II and their antennae in the thylakoid membrane plays an important role in this regulation.

In chloroplasts, the thylakoid membranes are present in two differ- ent arrays, as stacked and unstacked membranes. The outer surface of the unstacked membranes has free access to the stromal space; these mem- branes are called stromal lamellae (Fig. 3.40). In the stacked membranes, the neighboring thylakoid membranes are in direct contact with each other. These membrane stacks can be seen as grains (grana) in light microscopy and are therefore called granal lamellae.

ATP synthase and the PS I complex (including its light harvesting com- plexes, not further discussed here) are located either in the stromal lamellae or in the outer membrane region of the granal lamellae. Therefore, these proteins have free access to ADP and NADP^ in the stroma. The PS II complex, on the other hand, is primarily located in the granal lamellae. Peripheral LHC II subunits attached to the PS II complex (section 2.4) contain a protein chain protruding from the membrane, which can prob- ably interact with the LHC II subunit of the adjacent membrane and thus

cause tight membrane stacking. The cyt-b₆/f complexes are only present in stacked membranes. Since the proteins of PS I and F-ATP-synthase project into the stroma space, they do not fit into the space between the stacked membranes. Thus the PS II complexes in the stacked membranes are sepa- rated spatially from the PS I complexes in the unstacked membranes. It is assumed that this prevents an uncontrolled spillover of excitons from PS II to PS I.

However, the spatial separation of the two photosystems and thus the spillover of excitons from PS II to PS I can be regulated. For example, if the excitation of PS II is greater than that of PS I, plastohydroquinone accumulates, which cannot be oxidized rapidly enough via the cyt-b₆/f com- plex by PS I. Under these conditions, a protein kinase is activated, which phosphorylates the hydroxyl groups of threonine residues of peripheral LHC II subunits, causing a conformational change of the LHC protein. As a result of this, the affinity to PS II is decreased and the LHC II subunits dissociate from the PS II complexes. Furthermore, due to the changed con- formation, LHC II subunits can now bind to PS I, mediated by the H sub- unit of PS II. This LHC II-PS I complex purposely increases the spillover of excitons from LHC II to PS I. In this way the accumulation of reduced plastoquinone decreases the excitation of PS II and enhances the excitation of PS I. A protein phosphatase facilitates the reversal of this regulation. This regulatory process, which has been simplified here, enables an opti- mized distribution of the captured photons between the two photosystems, independent of the spectral quality of the absorbed light.

Excess light energy is eliminated as heat

Plants face the general problem that the energy of irradiated light can be much higher than the demand of photosynthetic metabolites such as NADPH and ATP. This is the case when very high light intensities are present and the metabolism cannot keep pace. Such a situation arises at low temperatures, when the metabolism is slowed down because of decreased enzyme activities (cold stress) or at high temperatures, when stomata close to prevent loss of water. Excess excitation of the photosystems could result in an excessive reduction of the components of the photosynthetic electron transport.

Very high excitation of photosystem II, recognized by the accumulation of plastohydroquinone, results in damage to the photosynthetic appara- tus, termed photoinhibition. A major cause of this damage is an overexcita- tion of the reaction center, by which chlorophyll molecules attain a triplet state, resulting in the formation of aggressive singlet oxygen (section 2.3). The damaging effect of triplet chlorophyll can be demonstrated by placing

a small amount of chlorophyll under the human skin, which after illumina- tion causes severe tissue damage. This photodynamic principle is utilized in medicine for the selective therapy of skin cancer. Carotenoids (e.g., –carotene, Fig. 2.9) are able to convert the triplet state of chlorophyll and the singlet state of oxygen to the corresponding ground states by forming a triplet carotenoid, which dissipates its energy as heat. In this way carotenoids have an important protective function. If under certain conditions this protective function of carotenoids is una- ble to cope with excessive excitation of PS II, the remaining singlet oxy- gen has a damaging effect on the PS II complex. The site of this damage could be the D₁ protein of the photosynthetic reaction center in PS II, which already under normal photosynthetic conditions experiences a high turnover (see section 3.6). When the rate of D₁-protein damage exceeds the rate of its resynthesis, the rate of photosynthesis is decreased, resulting in photoinhibition.

Plants have developed several mechanisms to protect the photosynthetic apparatus from light damage. One mechanism is chloroplast avoidance movement, in which chloroplasts move under high light conditions from the cell surface to the side walls of the cells. Another way is to dissipate the energy arising from an excess of excitons as heat. This process is termed nonphotochemical quenching of exciton energy. Although our knowledge of this quenching process is still incomplete, it is undisputed that zeaxan- thin plays an important role. Zeaxanthin causes the dissipation of exciton energy to heat by interacting with a chlorophyll-binding protein (CP 22) of photosystem II. Zeaxanthin is formed by the reduction of the diepoxide vio- laxanthin. The reduction proceeds with ascorbate as the reductant and the monoepoxide antheraxanthin is formed as an intermediate. Zeaxanthin can be reconverted to violaxanthin by epoxidation which requires NADPH and O₂ (Fig. 3.41). Formation of zeaxanthin by diepoxidase takes place on the luminal side of the thylakoid membrane at an optimum pH of 5.0, whereas the regeneration of violaxanthin by the epoxidase proceeding at the stro- mal side of the thylakoid membrane occurs at about pH 7.6. Therefore, the formation of zeaxanthin requires a high pH gradient across the thylakoid membrane. As discussed in connection with the Mehler reaction (section 3.9), a high pH gradient can be an indicator of the high excitation state of photosystem II. When there is too much excitation energy, an increased pH gradient initiates zeaxanthin synthesis, dissipating excess energy of the PS II complex as heat. This mechanism explains how under strong sunlight most plants convert 50% to 70% of all the absorbed photons to heat. The non-photochemical quenching of excitation energy is the primary way for plants to protect themselves from too much light energy. In comparison, the Mehler reaction (section 3.9) and photorespiration (section 7.7) under

Figure 3.35 A schemefor the Mehler reaction.Upon strong reduction offerredoxin, electrons aretransferred by the Mehlerreaction to oxygen andsuperoxide is formed. Theelimination of this highlyaggressive superoxideradical involves reactionscatalyzed by superoxidedismutase and ascorbateperoxidase

elker and Or: Soil Hydrology and Biophysics

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ps 2 – electron transport

Photosynthesis is an electron transport process

The previous chapter described how photons are captured by an antenna and conducted as excitons to the reaction centers. This chapter deals with the function of these reaction centers and describes how photon energy is converted to chemical energy to be utilized by the cell. As mentioned in Chapter 2, plant photosynthesis probably evolved from bacterial pho- tosynthesis, so that the basic mechanisms of the photosynthetic reactions are alike in bacteria and plants. Bacteria have proved to be very suitable objects for studying the principles of photosynthesis since their reaction centers are more simply structured than those of plants and they are more easily isolated. For this reason, first bacterial photosynthesis and then plant photosynthesis will be described.

The photosynthetic machinery is constructed from modules

The photosynthetic machinery of bacteria is constructed from defined com- plexes, which also appear as components of the photosynthetic machinery in plants. As will be described in Chapter 5, some of these complexes are also components of mitochondrial electron transport. These complexes can be thought of as modules that developed at an early stage of evolu- tion and have been combined in various ways for different purposes. For easier understanding, the functions of these modules in photosynthesis will be treated first as black boxes and a detailed description of their structure and function will be given later.

E °

Figure 3.1 Schematic presentation of the photosynthetic apparatus of purple bacteria. The energy of a captured exciton in the reaction center elevates an electron to a negative redox state. The electron is transferred to the ground state via an electron transport chain including the cytochrome-b/c₁ complex and cytochrome-c (the latter is not shown). Free energy of this process is conserved by formation of a proton potential which is used partly for synthesis of ATP and partly to enable an electron flow for the formation of NADH from electron donors such as H₂S.

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Oxidation state shows the total number of electrons which have been removed from an element (a positive oxidation state) or added to an element (a negative oxidation state) to get to its present state.

Oxidation involves an increase in oxidation state
Reduction involves a decrease in oxidation state

Section 6.1 – An Introduction to Oxidation Reduction Reactions (10:27)

Section 6.2 – Oxidation Numbers (22:09)

Section 6.4 – Voltaic Cells (24:17)

0.3: Electrochemical Potential

n a galvanic cell, current is produced when electrons flow externally through the circuit from the anode to the cathode because of a difference in potential energy between the two electrodes in the electrochemical cell. In the Zn/Cu system, the valence electrons in zinc have a substantially higher potential energy than the valence electrons in copper because of shielding of the s electrons of zinc by the electrons in filled d orbitals. Hence electrons flow spontaneously from zinc to copper(II) ions, forming zinc(II) ions and metallic copper. Just like water flowing spontaneously downhill, which can be made to do work by forcing a waterwheel, the flow of electrons from a higher potential energy to a lower one can also be harnessed to perform work….

Redox reactions can be balanced using the half-reaction method. The standard cell potential is a measure of the driving force for the reaction. \(E°_{cell} = E°_{cathode} − E°_{anode} \] The flow of electrons in an electrochemical cell depends on the identity of the reacting substances, the difference in the potential energy of their valence electrons, and their concentrations. The potential of the cell under standard conditions (1 M for solutions, 1 atm for gases, pure solids or liquids for other substances) and at a fixed temperature (25°C) is called the standard cell potential (E°_cell). Only the difference between the potentials of two electrodes can be measured. The potential of the standard hydrogen electrode (SHE) is defined as 0 V under standard conditions. The potential of a half-reaction measured against the SHE under standard conditions is called its standard electrode potential. By convention, all tabulated values of standard electrode potentials are listed as standard reduction potentials. The overall cell potential is the reduction potential of the reductive half-reaction minus the reduction potential of the oxidative half-reaction (E°_cell = E°_cathode − E°_anode). The standard cell potential is a measure of the driving force for a given redox reaction. If E°cell is positive, the reaction will occur spontaneously under standard conditions. If E°cell is negative, then the reaction is not spontaneous under standard conditions, although it will proceed spontaneously in the opposite direction.

Cyclic Electron Transport in Photosynthesis

Photosynthetic reaction center

The reaction center is in the thylakoid membrane. It transfers light energy to a dimer of chlorophyll pigment molecules near the periplasmic (or thylakoid lumen) side of the membrane. This dimer is called a special pair because of its fundamental role in photosynthesis. This special pair is slightly different in PSI and PSII reaction center. In PSII, it absorbs photons with a wavelength of 680 nm, and it is therefore called P680. In PSI, it absorbs photons at 700 nm, and it is called P700. In bacteria, the special pair is called P760, P840, P870, or P960. “P” here means pigment, and the number following it is the wavelength of light absorbed.

If an electron of the special pair in the reaction center becomes excited, it cannot transfer this energy to another pigment using resonance energy transfer. In normal circumstances, the electron should return to the ground state, but, because the reaction center is arranged so that a suitable electron acceptor is nearby, the excited electron can move from the initial molecule to the acceptor. This process results in the formation of a positive charge on the special pair (due to the loss of an electron) and a negative charge on the acceptor and is, hence, referred to as photoinduced charge separation. In other words, electrons in pigment molecules can exist at specific energy levels. Under normal circumstances, they exist at the lowest possible energy level they can. However, if there is enough energy to move them into the next energy level, they can absorb that energy and occupy that higher energy level. The light they absorb contains the necessary amount of energy needed to push them into the next level. Any light that does not have enough or has too much energy cannot be absorbed and is reflected. The electron in the higher energy level, however, does not want to be there; the electron is unstable and must return to its normal lower energy level. To do this, it must release the energy that has put it into the higher energy state to begin with. This can happen various ways. The extra energy can be converted into molecular motion and lost as heat. Some of the extra energy can be lost as heat energy, while the rest is lost as light. (This re-emission of light energy is called fluorescence.) The energy, but not the e- itself, can be passed onto another molecule. (This is called resonance.) The energy and the e- can be transferred to another molecule. Plant pigments usually utilize the last two of these reactions to convert the sun’s energy into their own.

This initial charge separation occurs in less than 10 picoseconds (10⁻¹¹ seconds). In their high-energy states, the special pigment and the acceptor could undergo charge recombination; that is, the electron on the acceptor could move back to neutralize the positive charge on the special pair. Its return to the special pair would waste a valuable high-energy electron and simply convert the absorbed light energy into heat. In the case of PSII, this backflow of electrons can produce reactive oxygen species leading to photoinhibition.^[1][2] Three factors in the structure of the reaction center work together to suppress charge recombination nearly completely.

Another electron acceptor is less than 10 Å away from the first acceptor, and so the electron is rapidly transferred farther away from the special pair.
An electron donor is less than 10 Å away from the special pair, and so the positive charge is neutralized by the transfer of another electron
The electron transfer back from the electron acceptor to the positively charged special pair is especially slow. The rate of an electron transfer reaction increases with its thermodynamic favorability up to a point and then decreases. The back transfer is so favourable that it takes place in the inverted region where electron-transfer rates become slower.^[1]

Thus, electron transfer proceeds efficiently from the first electron acceptor to the next, creating an electron transport chain that ends if it has reached NADPH.

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Purple bacteria have only one reaction center (Fig. 3.1). In this reac- tion center the energy of the absorbed photon excites an electron, which will be elevated to a negative redox state. The excited electron is transferred back to the ground state by an electron transport chain, called the cyto- chrome-b/c₁ complex, and the released energy is transformed to a chemical compound (NADH), which is then used for the synthesis of biomass (e.g., proteins and carbohydrates). Generation of energy is based on coupling the electron transport with the transport of protons across the membrane. In this way the energy of the excited electron is conserved as an electrochemical H^-potential across the membrane. The photosynthetic reaction cent- ers and the main components of the electron transport chain are always located in a membrane.

Via ATP-synthase the energy of the H^-potential is used to synthesize ATP from ADP and phosphate. Since the excited electrons in purple bacte- ria return to the ground state of the reaction center, this electron transport is called cyclic electron transport. In purple bacteria the proton gradient is also used to reduce NAD^ via an additional electron transport chain named the NADH dehydrogenase complex (Fig. 3.1). By consuming the energy of the H^-potential, electrons are transferred from a reduced compound (e.g., organic acids or hydrogen sulfide) to NAD^. The ATP and NADH formed by bacterial photosynthesis are used for the synthesis of organic matter; especially important is the synthesis of carbohydrates from CO₂ via the Calvin cycle (see Chapter 6).

The reaction center of green sulfur bacteria (Fig. 3.2) is homologous to that of purple bacteria, indicating that they have both evolved from a com- mon ancestor. ATP is also formed in green sulfur bacteria by cyclic electron transport. The electron transport chain (cytochrome-b/c₁ complex) and the ATP-synthase involved here are very similar to those in purple bacteria. However, in contrast to purple bacteria, green sulfur bacteria are able to synthesize NADH by a noncyclic electron transport process. In this case, the excited electrons are transferred to the ferredoxin-NAD-reductase complex, which reduces NAD^ to NADH. Since the excited electrons in this noncyc- lic pathway do not return to the ground state, an electron deficit remains in the reaction center and is replenished by electron donors such as H₂S, ulti- mately being oxidized to sulfate.

Cyanobacteria and plants use water as an electron donor in photosyn thesis (Fig. 3.3). As oxygen is liberated, this process is called oxygenic pho- tosynthesis. Two photosystems designated II and I are arranged in tandem. The machinery of oxygenic photosynthesis is built by modules that have already been described in bacterial photosynthesis. The structure of the reaction center of photosystem II corresponds to that of the reaction center of purple bacteria, and that of photosystem I corresponds to the reaction center of green sulfur bacteria. The enzymes ATP synthase and ferredoxin- NADP-reductase are very similar to those of photosynthetic bacteria. The

Figure 3.2 Schematic presentation of the photosynthetic apparatus in green sulfur bacteria. In contrast to the scheme in Figure 3.1, part of the electrons elevated to a negative redox state is transferred via an electron transport chain (ferredoxin- NAD reductase) to NAD, yielding NADH. The electron deficit arising in the reaction center is compensated by electron donors such as H2S (see also Fig. 3.1).

Figure 3.3 Schematic presentation of the photosynthetic apparatus of cyanobacteria and plants. The two sequentially arranged reaction centers correspond in their function to the photosynthetic reaction centers of purple bacteria and green sulfur bacteria (shown in Figs. 3.1 and 3.2).

electron transport chain of the cytochrome-b₆/f complex has the same basic structure as the cytochrome-b/c₁ complex in bacteria.

Four excitons are required in oxygenic photosynthesis to split one mol- ecule of water:

H2O  NADP^  4 excitons → 1/2 O2  NADPH  H^

In this noncyclic electron transport, electrons are transferred to NADP^ and protons are transported across the membrane to generate a proton gradient that drives the synthesis of ATP. Thus, for each mol of NADPH formed by oxygenic photosynthesis, about 1.5 molecules of ATP are gener- ated simultaneously (section 4.4). Most of this ATP and NADPH are used for CO₂ and nitrate assimilation to synthesize carbohydrates and amino acids. Oxygenic photosynthesis in plants takes place in the chloroplasts, a cell organelle of the plastid family (section 1.3).

Vides

Redox Reactions

o-Oxidation and reduction review from biological point-of-view | Biomolecules | MCAT | Khan Academy

Oxidation and reduction in cellular respiration | Biology | Khan Academy

A reductant and an oxidant are formed during photosynthesis

In the 1920s Otto Warburg (Berlin) postulated that the energy of light is transferred to CO₂ and that the CO₂, activated in this way, reacts with water to form a carbohydrate, accompanied by the release of oxygen. According to this hypothesis, the oxygen released by photosynthesis was derived from the CO₂. In 1931 this hypothesis was opposed by Cornelis van Niel (USA) by postulating that during photosynthesis a reductant is formed, which then reacts with CO₂. The so-called van Niel equation describes photosynthesis in the following way: He proposed that a compound H₂A is split by light energy into a reduc- ing compound (2H) and an oxidizing compound (A). For oxygenic photo- synthesis of cyanobacteria or plants, it can be rewritten as:

In this equation the oxygen released during photosynthesis is derived from water. In 1937 Robert Hill (Cambridge, UK) proved that a reductant is actu- ally formed in the course of photosynthesis. He was the first to succeed in isolating chloroplasts with photosynthetic activity, which, however, had no intact envelope membranes and consisted only of thylakoid membranes. When these chloroplasts were illuminated in the presence of Fe^3 com- pounds (initially ferrioxalate, later ferricyanide ([Fe (CN)₆]^3)), Robert Hill observed an evolution of oxygen accompanied by the reduction of the Fe³^-compounds to the Fe²^ form.

This “Hill reaction” proved that the photochemical splitting of water can be separated from the reduction of the CO₂. Therefore the complete reac- tion of photosynthetic CO₂ assimilation can be divided into two reactions:

The so-called light reaction, in which water is split by photon energy to yield reductive power (NADPH) and chemical energy (ATP); and

the so-called dark reaction (Chapter 6), in which CO₂ is assimilated at the expense of the reductive power and of ATP.

In 1952 the Dutchman Louis Duysens made a very important observa- tion that helped explain the mechanism of photosynthesis. When illumi- nating isolated membranes of the purple bacterium Rhodospirillum rubrum with short light pulses, he found a decrease in light absorption at 890 nm, which was immediately reversed when the bacteria were darkened again. The same “bleaching” effect was found at 870 nm in the purple bacte- rium Rhodobacter sphaeroides. Later, Bessil Kok (USA) and Horst Witt (Germany) also found similar pigment bleaching at 700 nm and 680 nm in chloroplasts. This bleaching was attributed to the primary reaction of pho- tosynthesis, and the corresponding pigments of the reaction centers were named P₈₇₀ (Rb. sphaeroides) and P₆₈₀ and P₇₀₀ (chloroplasts). When an oxidant (e.g., [Fe(CN)₆]^3) was added, this bleaching effect could also be achieved in the dark. These results indicated that these absorption changes of the pigments were due to a redox reaction. This was the first indication that chlorophyll can be oxidized. Electron spin resonance measurements revealed that radicals are formed during this “bleaching.” “Bleaching” could also be observed at the very low temperature of 1 K. This showed that in the electron transfer leading to the formation of radicals, the reac- tion partners are located so close to each other that thermal oscillation of the reaction partners (normally the precondition for a chemical reaction) is not required for this redox reaction. Spectroscopic measurements indicated that the reaction partner of this primary redox reaction are two closely adjacent chlorophyll molecules arranged as a pair, called a “special pair.”

The basic structure of a photosynthetic reaction center has been resolved by X-ray structure analysis

The reaction centers of purple bacteria proved to be especially suitable objects for explaining the structure and function of the photosynthetic machinery. It was a great step forward when in 1970 Roderick Clayton (USA) developed a method for isolating reaction centers from purple bacte- ria. Analysis of the components of the reaction centers of the different purple bacteria (shown in Table 3.1 for the reaction center of Rhodobacter sphaer- oides as an example) revealed that the reaction centers had the same basic structure in all the purple bacteria investigated. The minimum structure

consists of the three subunits L, M, and H (light, medium, and heavy). Subunits L and M are peptides with a similar amino acid sequence. They are homologous. The reaction center of Rb. sphaeroides contains four bac- teriochlorophyll-a (Bchl-a, Fig. 3.4) and two bacteriopheophytin-a (Bphe- a). Pheophytins differ from chlorophylls in that they lack magnesium as the central atom. In addition, the reaction center contains an iron atom that is not part of a heme. It is therefore called a non-heme iron. Furthermore, the reaction center is comprised of two molecules of ubiquinone (Fig. 3.5), which are designated as Q_A and Q_B. Q_A is tightly bound to the reaction center, whereas Q_B is only loosely associated with it.

Figure 3.5 Ubiquinone. The long isoprenoid side chain mediates the lipophilic character and membrane anchorage.

X-ray structure analysis of the photosynthetic reaction center

If ordered crystals can be prepared from a protein, it is possible to analyze the spherical structure of the protein molecule by X-ray structure analysis. In this method a protein crystal is exposed to X-ray irradiation. The elec- trons of the atoms in the molecule cause a scattering of X-rays. Diffraction is observed when the irradiation passes through a regular repeating struc- ture. The corresponding diffraction pattern, consisting of many single reflections, is measured by an X-ray film positioned behind the crystal or by an alternative detector. The principle is demonstrated in Figure 3.6. To obtain as many reflections as possible, the crystal, mounted in a capillary, is rotated. From a few dozen to up to several hundred exposures are required for one set of data, depending on the form of the crystal and the size of the crystal lattice. To evaluate a new protein structure, several sets of data are required in which the protein has been changed by the incorporation or binding of a heavy metal ion. With the help of elaborate computer pro- grams, it is possible to reconstruct the spherical structure of the exposed protein molecules by applying the rules for scattering X-rays by atoms of various electron densities. slowly and the diffraction pattern is monitored on an X-ray film. Nowadays much more sensitive detector systems (image platers) are used instead of films. The diffraction pattern shown was obtained by the structural analysis of the reaction center of

Rb. sphaeroides. (Courtesy of H. Michel, Frankfurt.)

X-ray structure analysis requires a highly technical expenditure and is very time-consuming, but the actual limiting factor in the elucidation of a spherical structure is usually the preparation of suitable single crystals. Until 1980 it was thought to be impossible to prepare crystals suitable for X-ray structure analysis from hydrophobic membrane proteins. The appli- cation of the detergent N,N-dimethyldodecylamine-N-oxide (Fig. 3.7) was a great step forward in helping to solve this problem. This detergent forms water-soluble protein-detergent micelles with membrane proteins. With the addition of ammonium sulfate or polyethylene glycol is was then pos- sible to crystallize membrane proteins. The micelles form a regular lattice in these crystals (Fig. 3.8). The protein in the crystal remains in its native state since the hydrophobic regions of the membrane protein, which nor- mally border on the hydrophobic membrane, are covered by the hydropho- bic chains of the detergent.

Using this procedure, Hartmut Michel (Munich) succeeded in obtaining crystals from the reaction center of the purple bacterium Rhodopseudomonas viridis and, together with his colleagues Johann Deisenhofer and Robert Huber, performed an X-ray structure analysis of these crystals. The immense amount of time invested in these investigations is illustrated by the fact that the evaluation of the stored data sets alone took two and a half years (nowa- days modern computer programs would do it very much faster). The X-ray structure analysis of a photosynthetic reaction center successfully elucidated for the first time the three-dimensional structure of a membrane protein. For this work, Michel, Deisenhofer, and Huber were awarded the Nobel Prize in Chemistry in 1988. Using the same method, the reaction center of Rb. sphaeroides was analyzed and it turned out that the basic structures of the two reaction centers are astonishingly similar.

Figure 3.6 Schematic presentation of X-ray structural analysis of a protein crystal. A capillary containing the crystal rotates slowly and the diffraction pattern is monitored on an X-ray film. Nowadays muc more sensitive detector systems (image platers) are used instead of films. The diffraction pattern shown was obtained by the structural analysis of the reaction center of Rb. sphaeroides. (Courtesy of H. Michel, Frankfurt.)

The reaction center of Rhodopseudomonas viridis has a symmetric structure

Figure 3.9 shows the three-dimensional structure of the reaction center of the purple bacterium Rhodopseudomonas viridis. The molecule has a cylin- drical shape and is about 8 nm long. The homologous subunits L (red) and

Figure 3.8 A micelle is formed after solubilization of a membrane protein with detergent. The hydrophobic region of the membrane proteins, the membrane lipids, and the detergent are shown in black and the hydrophilic regions in red. Crystal structures can be formed by association of the hydrophilic regions of the detergent micelle.

M (black) are arranged symmetrically and enclose the chlorophyll and phe- ophytin molecules. The H subunit is attached like a lid to the lower part of the cylinder. In the same projection as in Figure 3.9, Figure 3.10 shows the location of the chromophores in the protein molecule. All the chromophores are positioned as pairs divided by a symmetry axis. Two Bchl-a molecules (D_M, D_L) can be recognized in the upper part of the structure. The two tetrapy- rrole rings are so close (0.3 nm) that their orbitals overlap in the excited state. This confirmed the actual existence of the “special pair” of chloro- phyll molecules, postulated earlier from spectroscopic investigations, as the

site of the primary redox process of photosynthesis. The chromophores are arranged underneath the chlorophyll pair in two nearly identical branches, both comprised of a Bchl-a (B_A, B_B) monomer and a bacteriopheophytin (_A, _B). Whereas the chlorophyll pair (D_M, D_L) is bound by both sub- units L and M, the chlorophyll B_A and the pheophytin _A are associated with subunit L, and B_B and _B with subunit M. The quinone ring of Q_A is bound via hydrogen bonds and hydrophobic interaction to subunit M, whereas the loosely associated Q_B is bound to subunit L.

playlist–

Photosynthesis

Photosystem II Function: The P680 Reaction Center

How does a reaction center function?

The analysis of the structure and extensive kinetic investigations allowed a detailed description of the function of the bacterial reaction center. The kinetic investigations included measurements by absorption and fluores- cence spectroscopy after light flashes in the range of less than 10^13 s, as well as measurements of nuclear spin and electron spin resonance. Although the reaction center shows a symmetry with two almost identical branches of chromophores, electron transfer proceeds only along the right branch (the L side, Fig. 3.10). The chlorophyll monomer (B_B) on the M side is in close contact with a carotenoid molecule, which abolishes a harmful triplet state of chlorophylls in the reaction center (sections 2.3 and 3.7). The function of the pheophytin (_B) on the M side and of the non-heme iron is not yet fully understood.

Figure 3.11 presents a scheme of the reaction center with the reaction partners arranged according to their electrochemical potential. The exciton of the primary reaction is provided by the antenna (section 2.4) which then excites the chlorophyll pair. This primary excitation state has only a very short half-life time, then a charge separation occurs, and, as a result of the large potential difference, an electron is removed within picoseconds to reduce bacteriopheophytin (Bphe).

The electron is probably transferred first to the Bchl-monomer (B_A) and then to the pheophytin molecule (_A). The second electron transfer proceeds with a half-time of 0.9 picoseconds, about four times as fast as the elec- tron transfer to B_A. The pheophytin radical has a tendency to return to the ground state by a return of the translocated electron to the Bchl-monomer (B_A). To prevent this, within 200 picoseconds a high potential difference withdraws the electron from the pheophytin radical to a quinone (Q_A) (Fig. 3.11). The semiquinone radical thus formed, in response to a further poten- tial difference, transfers its electron to the loosely bound ubiquinone Q_B. After a second electron transfer, first ubisemiquinone and then ubihydroqui- none are formed (Fig. 3.12). In contrast to the very labile radical intermedi- ates of the pathway, ubihydroquinone is a stable reductant. However, this stability has its price. For the formation of ubihydroquinone as a first sta- ble product from the primary excitation state of the chlorophyll, more than half of the exciton energy is dissipated as heat.

Ubiquinone (Fig. 3.5) contains a hydrophobic isoprenoid side chain, by which it is soluble in the lipid phase of the photosynthetic membrane. The same function of an isoprenoid side chain has already been discussed in the case of chlorophyll (section 2.2). In contrast to chlorophyll, pheophytin, and Q_A, which are all tightly bound to proteins, ubihydroquinone Q_B is only loosely associated with the reaction center and can be exchanged by another ubiquinone. Ubihydroquinone remains in the membrane phase, is able to diffuse rapidly along the membrane, and functions as a transport metabolite for reducing equivalents in the membrane phase. It feeds the electrons into the cytochrome-b/c₁ complex, also located in the membrane. The electrons are then transferred back to the reaction center through the cytochrome-b/c₁

complex and via cytochrome-c. Energy is conserved during this electron transport as a proton potential (section 4.1), which is used for ATP-synthe- sis. The structure and mechanism of the cytochrome-b/c₁ complex and of ATP-synthase will be described in section 3.7 and Chapter 4, respectively.

In summary, the cyclic electron transport of the purple bacteria resembles a simple electric circuit (Fig. 3.13). The two pairs of chlorophyll and pheophy- tin, between which an electron is transferred by light energy, may be regarded as the two plates of a capacitor between which a voltage is generated, driv- ing a flux of electrons, a current. Voltage drops via a resistor and a large amount of the electron energy is dissipated as heat. This resistor functions

as an electron trap, and withdraws the electrons rapidly from the capacitor. A generator utilizes the remaining voltage to produce chemical energy.

Figure 3.11 Schematic presentation of cyclic electron transport in photosynthesis of Rb. sphaeroides. The excited state symbolized by a star results in a charge separation; an electron is transferred via pheophytin, the quinones QA, QB, and the cyt-b/c complex to the positively charged chlorophyll radical. Q: quinone, Q•  : semiquinone radical, QH2: hydroquinone.

Figure 3.12 Reduction ofa quinone by one electronresults in a semiquinoneradical and furtherreduction to hydroquinone.Q: quinone, Q•  :semiquinone radical, QH2:hydroquinone.ExcitonGeneratorCyt-b/c1complexElectrontrap+ –Chlorophyll dimerHeatchemicalworkATP

Figure 3.13 Cyclic electrontransport of photosynthesisdrawn as an electricalcircuit.

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ps 1 – light harvesting

The use of energy from sunlight by photosynthesis is the basis of life on earth

Chlorophyll is the main photosynthetic pigment

Pigments capture energy from sunlight

The energy content of light depends on its wavelength

In Berlin at the beginning of the twentieth century Max Planck and Albert Einstein, two Nobel Prize winners, carried outthe epoch-making studies proving

that light has a dual nature. It can be regarded as an electromagnetic wave as well as an emission of particles, which are termed light quanta or photons.

The energy of the photon is proportional to its frequency v:

where h is the Planck constant (6.6 · 10^34 J s) and c the velocity of the light (3 · 10⁸ m s^1).  is the wavelength of light.

The mole (abbreviated to mol) is used as a chemical measure for the amount of molecules and the amount of photons corresponding to 6 · 10²³ molecules or photons (Avogadro number N_A). The energy of one mol photons amounts to:

In order to utilize the energy of a photon in a thermodynamic sense, this energy must be at least as high as the Gibbs free energy of the photochemi- cal reaction involved. (In fact much energy is lost during energy conversion (section 3.4), with the consequence that the energy of the photon must be higher than the Gibbs free energy of the corresponding reaction.) We can equate the Gibbs free energy G with the energy of the absorbed light:

The introduction of numerical values of the constants h, c, and N_A yields:

The human eye perceives only the small range between about 400 and 700 nm of the broad spectrum of electromagnetic waves (Fig. 2.2). The light in this range, where the intensity of solar radiation is especially high, is uti- lized in plant photosynthesis. Bacterial photosynthesis, however, is able to utilize light in the infrared range.

According to equation 2.3 the energy of irradiated light is inversely proportional to the wavelength. Table 2.1 shows the light energy per mol photons for light of different colors. Consequently, violet light has an energy of about 300 kJ/mol photons. Dark blue light, with the highest wavelength (700 nm) that can still be utilized by plant photosynthesis, contains 170 kJ/ mol photons. This is only about half the energy content of violet light.

In photosynthesis of a green plant, light is collected primarily by chlorophylls, pigments that absorb light at a wavelength below 480 nm and between 550 and 700 nm (Fig. 2.3). When white sunlight falls on a chlorophyll layer, the green light with a wavelength between 480 and 550 nm is not absorbed, but is reflected. This is why plant chlorophylls and whole leaves appear green.

Experiments carried out between 1905 and 1913 in Zurich and Berlin by Richard Willstätter and his collaborators led to the discovery of the structural formula of the green leaf pigment chlorophyll, a milestone in the history of chemistry. This discovery made such an impact that Richard Willstätter was awarded the Nobel Prize in Chemistry as early as 1915. There are different classes of chlorophylls. Figure 2.4 shows the structural formulas of chlorophyll-a and chlorophyll-b (chl-a, chl-b). The basic struc- ture is a ring made of four pyrroles, a tetrapyrrole, which is also named porphyrin. Mg^ is present in the center of the ring as the central atom. Mg^ is covalently bound with two N atoms and coordinately bound to the other two atoms of the tetrapyrrole ring. A cyclopentanone is attached to ring c. At ring d a propionic acid group forms an ester with the alcohol phytol. Phytol consists of a long branched hydrocarbon chain with one C-C double bond. It is derived from an isoprenoid, formed from four isoprene units (section 17.7). This long hydrophobic hydrocarbon tail renders the chlorophyll highly soluble in lipids and therefore promotes its presence in the membrane phase. Chlorophyll always occurs bound to proteins. Chl-b contains a formyl residue in ring b instead of the methyl residue as in chl-a. This small difference has a large influence on light absorption. Figure 2.3 shows that the absorption spectra of chl-a and chl-b differ markedly.

In plants, the ratio chl-a to chl-b is about three to one. Only chl-a is a constituent of the photosynthetic reaction centers (sections 3.6 and 3.8) and therefore it can be regarded as the central photosynthesis pigment. In a wide range of the visible spectrum, however, chl-a does not absorb light (Fig. 2.3). This non-absorbing region is named the “green window.” The absorption gap is narrowed by the light absorption of chl-b, with its first maximum at a higher wavelength than chl-a and the second maximum at a lower wavelength. As shown in section 2.4, the light energy absorbed by chl-b can be transferred very efficiently to chl-a. In this way, chl-b enhances the plant’s efficiency for utilizing sunlight energy.

The structure of chlorophylls has remained remarkably constant during the course of evolution. Purple bacteria, probably formed more than 3 billion years ago, contain as photosynthetic pigment a bacteriochlorophyll-a, which differs from the chl-a shown in Fig. 2.4 only by the alteration of one side chain and by the lack of one double bond. This, however, influences light absorption; both absorption maxima are shifted outwards and the non-absorbing spectral region in the middle is broadened. This shift allows purple bacteria to utilize light in the infrared region.

The tetrapyrrole ring not only is a constituent of chlorophyll but also has attained a variety of other functions during evolution. It is involved in methane formation by bacteria with Ni^ as the central atom. With Co^ it forms cobalamin (vitamin B₁₂), which participates as a cofactor in reac- tions in which hydrogen and organic groups change their position. With Fe^ instead of Mg^ as the central atom, the tetrapyrrole ring forms the basic structure of hemes (Fig. 3.24), which as cytochromes function as redox carriers in electron transport processes (sections 3.7 and 5.5) and as myoglobin or hemoglobin stores or transports oxygen in aerobic organ- isms. The tetrapyrrole ring in animal hemoglobin differs only slightly from the tetrapyrrole ring of chl-a (Fig. 2.4).

It seems remarkable that a substance that attained a certain function during evolution is being utilized after only minor changes for completely different functions. The reason for this functional variability is that the reactivity of compounds such as chlorophyll or heme is governed to a great extent by the proteins to which they are bound.

Chlorophyll molecules are bound to chlorophyll-binding proteins. In a complex with proteins the absorption spectrum of the bound chlorophyll differs considerably from the absorption spectrum of the free chlorophyll. The same applies for other light-absorbing compounds, such as carotenoids, xanthophylls, and phycobilins, which also occur bound to proteins. These complexes will be discussed in the following sections. For better discrimi- nation in this text book, free absorbing compounds are called chromophore (Greek, carrier of color) and the chromophore-protein complexes are called pigments. Pigments are further characterized by the wavelength of their absorption maximum. Chlorophyll-a₇₀₀ describes a pigment of protein-chl-a complex with an absorption maximum of 700 nm. Another common desig- nation is P₇₀₀; this nomination leaves the nature of the chromophore open.

Light absorption excites the chlorophyll molecule

What happens when a chromophore absorbs a photon? When a photon with a certain wavelength hits a chromophore molecule that absorbs light

of this wavelength, the energy of the photon excites electrons and trans- fers them to a higher energy level. This occurs as an “all or nothing” proc- ess. According to the principle of energy conservation expressed by the first law of thermodynamics, the energy of the chromophore is increased by the energy of the photon, which results in an excited state of the chromo- phore molecule. The energy is absorbed only in discrete quanta, resulting in discrete excitation states. The energy required to excite a chromophore molecule depends on the chromophore structure. A general property of chromophores is that they contain many conjugated double bonds, 10 in the case of the tetrapyrrole ring of chl-a. These double bonds are delocalized. Figure 2.5 shows two possible resonance forms.

After absorption of energy, an electron of the conjugated system is ele- vated to a higher orbit. This excitation state is termed singlet. Figure 2.6 shows a scheme of the excitation process. As a rule, the higher the number of double bonds in the conjugated system, the lower the amount of energy required to produce a first singlet state. For the excitation of chlorophyll, dark red light is sufficient, whereas butadiene, with only two conjugated double bonds, requires energy-rich ultraviolet light for excitation. The light absorption of the conjugated system of the tetrapyrrole ring is influenced by the side chains. Thus, the differences in the absorption maxima of chl-a and chl-b mentioned previously can be explained by an electron attracting effect of the carbonyl side chain in ring b of chl-b (Fig. 2.5).

The spectra of chl-a and chl-b (Fig. 2.3) each have two main absorption maxima, showing that each chlorophyll has two main excitation states. In addition, chlorophylls have minor absorption maxima, which for the sake of simplicity will not be discussed here. The two main excitation states of chlo- rophyll are known as the first and second singlet (Fig. 2.6). The absorption

maxima in the spectra are relatively broad. At a higher resolution the spec- tra can be shown to consist of many separate absorption lines. This fine structure of the absorption spectra is due to chlorophyll molecules that are in the ground and in the singlet states as well in rotation and vibration. In the energy scheme the various rotation and vibration energy levels are drawn as fine lines and the corresponding ground states as solid lines (Fig. 2.6).

The energy levels of the various rotation and vibration states of the ground state overlap with the lowest energy levels of the first singlet. Analogously, the energy levels of the first and the second singlet also over- lap. If a chlorophyll molecule absorbs light in the region of its absorption maximum (blue light), one of its electrons is elevated to the second singlet state. This second singlet state with a half-life of only 10¹² s is too unsta- ble to use its energy for chemical work. The excited molecules lose energy as heat by rotations and vibrations until the first singlet state is reached. This first singlet state can also be attained by absorption of a photon of red light, which contains less energy. The first singlet state is much more stable than the second one; its half-life time is 4 · 10⁹ s.

The return of the chlorophyll molecule from the first singlet state to the ground state can proceed in different ways:

The most important path for the conversion of the energy released upon the return of the first singlet state to the ground state is its utiliza- tion for chemical work. The chlorophyll molecule transfers the excited electron from the first singlet state to an electron acceptor and a pos- itively charged chlorophyll radical chl^ remains. This is possible since the excited electron is bound less strongly to the chromophore molecule than in the ground state. Section 3.5 describes in detail how the elec- tron can be transferred back from the acceptor to the chl^ radical via an electron transport chain. When the chlorophyll molecule returns to the ground state, the free energy derived from this process is conserved for chemical work. As an alternative, the electron deficit in the chl^ radical may be replenished by another electron donor (e.g., water (section 3.6)).

The excited chlorophyll can return to the ground state by releasing exci- tation energy as light; this emitted light is named fluorescence. Due to vibrations and rotations, part of the excitation energy is usually lost as heat, with the result that the fluorescence light has less energy (corre- sponding to a longer wavelength) than the energy of the excitation light, which was required for attaining the first singlet state (Fig. 2.7).

It is also possible that the return from the first singlet to the ground state proceeds in a stepwise fashion via the various levels of vibration and rotation energy, by which the energy difference is completely con- verted into heat.

By releasing part of the excitation energy as heat, the chlorophyll mol- ecule can attain a lower energy excitation state, called the first triplet state. This triplet state cannot be reached directly from the ground state by excitation, since the spin of the excited electrons has been reversed. Since the probability of a reversal spin is low, the triplet state does not occur frequently. In the case of a very high excitation, however, some of the electrons of the chlorophyll molecules can reach this state. By emitting so-called phosphorescent light, the molecule can return from the triplet state to the ground state. Phosphorescent light is lower in energy than the light required to attain the first singlet state. The return from the triplet state to the ground state requires a reversal of the electron spin. As this is rather improbable, the triplet state, in comparison to the first singlet state, has a relatively long half-life time (10^4 to 10^2 s). The triplet state of the chlorophyll has no function in photosynthesis per se. In its triplet state, however, the chlorophyll can excite oxygen to a sin- glet state, whereby the oxygen becomes very reactive (reactive oxygen species, ROS, section 5.7) with a damaging effect on cell constituents. Section 3.10 describes how the plant manages to protect itself from the harmful singlet oxygen.

The return to the ground state can be coupled with the excitation of a neighboring chromophore molecule. This transfer is important for the function of the antennae and will be described in the following section.

Figure 2.2 Spectrum of the electromagnetic radiation. The enlargement in red illustrates the visible spectrum.

Figure 2.3 Absorption spectrum of chlorophyll-a (chl-a), chlorophyll-b (chl-b) and of the xanthophyll lutein dissolved in acetone. The intensity of the sun’s radiation at different wavelengths is given as a comparison

Video- Excitation of Chlorophyll by Light

Video- The Chloroplast

Table 2.1: The energy content and the electrochemical potential difference of photons of different wavelengths

Wavelengths (nm)	Light color	Energy content kJ/mol photons	E e volt
700	Red	170	1.76
650	Bright red	183	1.90
600	Yellow	199	2.06
500	Blue green	238	2.47
440	Blue	271	2.80
400	Violet	298	3.09

.4 Structural formula of chlorophyll-a. In chlorophyll-b the methyl group in ring b is replaced by a formyl group (A). The phytol side chain in red gives chlorophyll a lipid character.

Figure 2.4 Structural formula of chlorophyll-a. In chlorophyll-b the methyl group in ring b is replaced by a formyl group (A). The phytol side chain in red gives chlorophyll a lipid character.

Video- Emission and Absorption Spectra

Video- Light Absorption, Reflection, and Transmission

Video- Light-Harvesting: Photosynthetic Pigments and Exciton Transfer (Playlist photosynthesis)

Video- Light-Harvesting: The Antenna Complex

BREAK

Resonance structures of chlorophyll-a. In the region marked red, the double bonds are not localized; the  electrons are distributed over the entire conjugated system. The formyl residue of chlorophyll-b attracts electrons and thus affects the  electrons of the conjugated system.

Figure 2.5 Resonance structures of chlorophyll-a. In the region marked red, the double bonds are not localized; the  electrons are distributed over the entire conjugated system. The formyl residue of chlorophyll-b attracts electrons and thus affects the  electrons of the conjugated system.

Schematic presentation of the excitation states of chlorophyll-a and their return to the ground state. The released excitation energy is converted

Figure 2.6 Schematic presentation of the excitation states of chlorophyll-a and their return to the ground state. The released excitation energy is converted

An antenna is required to capture light

In order to excite a photosynthetic reaction center, a photon with defined energy content has to react with a chlorophyll molecule in the reaction center. The probability is very low that a photon not only has the proper energy, but also hits the pigment exactly at the site of the chlorophyll mol- ecule. Therefore efficient photosynthesis is possible only when the energy of photons of various wavelengths is captured over a certain surface by a so-called antenna (Fig. 2.8). Similarly, radio and television sets could not work without an antenna.

The antennae of plants consist of a large number of protein-bound chlorophyll molecules that absorb photons and transfer their energy to the reaction center. Only a few thousandths of the chlorophyll molecules in the leaf are constituents of the actual reaction centers; the remainder are contained in the antennae. Observations made as early as 1932 by Robert Emerson and William Arnold in the United States indicated that the large majority of chlorophyll molecules are not part of the reaction centers. The two researchers illuminated a suspension of the green alga Chlorella with light pulses of 10 s duration, interrupted by dark intervals of 20 ms. Evolution of oxygen was used as a measure for photosynthesis. The light pulses were made so short that chlorophyll could undergo only one photo- synthetic excitation cycle and a high light intensity was chosen in order to achieve maximum oxygen evolution. Apparently the photosynthetic appa- ratus was thus saturated with photons. Analysis of the chlorophyll content of the algae suspension showed that under saturating conditions only one molecule of O2 was formed per 2,400 chlorophyll molecules.

In the following years Robert Emerson refined these experiments and

was able to show when pulses were applied at very low light intensity, the amount of oxygen formed increased proportionally with the light intensity. From this it was calculated that the release of one molecule of oxygen had a minimum quantum requirement of about eight photons. These results set- tled a long scientific dispute with Otto Warburg, who had concluded from his experiments that only four photons are required for the evolution of one molecule of O2. Later it was recognized that each of the two reaction cent- ers requires four photons for the formation of O2. Moreover, the results of Emerson and Arnold allowed the calculation that about 300 chlorophyll molecules are associated with one reaction center. These are constituents of the antennae. The antennae contain additional accessory pigments to utilize those photons where the wavelength corresponds to the “green window” between the absorption maxima of the chlorophylls. In higher plants these pigments are carotenoids, mainly xanthophylls, including lutein and the related vio- laxanthin as well as carotenes such as -carotene to name the major com- pound (Fig. 2.9). Moreover, an important function of these carotenoids in the antennae is to prevent the formation of the harmful triplet state of the chlorophylls (section 3.10). Important constituents of the antennae in cyanobacteria are phycobilins, which will be discussed at the end of this chapter.

How is the excitation energy of the photons captured in the antennae and transferred to the reaction centers?

The transfer of energy in the antennae via electron transport from chromo- phore to chromophore in a sequence of redox processes, as in the elec- tron transport chains of photosynthesis or of mitochondrial respiration (Chapters 3 and 5), could be excluded, since such an electron transport would need considerable activation energy. This is not the case, since a flux of excitation energy can be measured in the antennae at temperatures as low as 1 K. At these low temperatures light absorption and fluorescence still occur, whereas chemical processes catalyzed by enzymes are completely inactive. Under these conditions the energy transfer in the antennae pro- ceeds according to a mechanism that is related to those of light absorption and fluorescence.

When chromophores are positioned very close to each other, the quan- tum energy of an irradiated photon is transferred from one chromophore to the next. One quantum of light energy is named a photon, one quantum of excitation energy transferred from one molecule to the next is termed an exciton. A prerequisite for the transfer of excitons is a specific position- ing of the chromophores. This is arranged by proteins, and therefore the chromophores of the antennae always occur as protein complexes.

The antennae of plants consist of an inner part and an outer part (Fig. 2.10). The outer antenna, formed by the light harvesting complexes (LHCs), col- lects the light. The inner antenna, consisting of the core complexes, is an integral constituent of the reaction centers; it also collects light and con- ducts the excitons that were collected in the outer antenna to the photosyn- thetic reaction centers.

The LHCs are composed of polypeptides, which bind chl-a, chl-b, xan- thophylls, and carotenes. These proteins, termed LHC polypeptides, are encoded in the nucleus. A plant contains many different LHC polypeptides. In a tomato, for instance, at least 19 different genes for LHC polypeptides have been found, which are very similar to each other and are members of a multigene family. They are homologous, as they have all evolved from a common ancestor.

Plants contain two reaction centers, which are arranged in sequence: a reaction center of photosystem II (PS II), which has an absorption maxi- mum at 680 nm, and a photosystem I (PS I) with an absorption maximum at 700 nm. The function of these reaction centers will be described in sec- tions 3.6 and 3.8. Both photosystems are composed of different LHCs.

The function of an antenna is illustrated by the antenna of photosystem II

The antenna of the PS II reaction center contains primarily four LHCs termed LHC-IIa–d. The main component is LHC-IIb; it represents 67% of

Photons are collected by an antenna and their energy is transferred to the reaction center. In this scheme the squares represent chlorophyll molecules. The excitons conducted to the reaction center cause a charge separation (section

Figure 2.8 Photons are collected by an antenna and their energy is transferred to the reaction center. In this scheme the squares represent chlorophyll molecules. The excitons conducted to the reaction center cause a charge separation (section

Figure 2.9 Structural formula of -carotene and of two xanthophylls (lutein

vcideo- 3 Major Classes of Pigments in Photosynthesis

video-Photosynthetic Electron Transport

Oxidation and reduction in cellular respiration | Biology | Khan Academy

Photosynthesis and Respiration

Photosynthesis

Z scheme/light reaction of photosynthesis with reduction potential

Redox Reactions and the light reactions of photosynthesis– 5.7 – Biol 189

Figure 2.10 Schematic presentation of a higher plant antenna

the light harvesting complexes i, phycobilisome

Figure 2.13 Schematic presentation of the light harvesting complexes in the antenna of photosystem II in a plant viewed from above (after Thornber); (a) LHC-IIa, (b) LHC-IIb. The inner antenna complexes are linked to the core complex by LHC-IIa and LHC-IIc (c) monomers. The function of the LHC- IId (d) and LHC-IIe (e) monomers is not entirely known

Figure 2.14 Schematic presentation of a side view of the structure of a phycobilisome. Each of the units shown consists of three - and three - subunits. (After Bryanth.)

Light-Harvesting: The Antenna Complex

phycobilisome : Energy and Electron Transfer in Photosynthetic Organisms,

Photosynthesis: What are Photosystems? Resonance, Reaction Centers and Antenna Complex?

Fate of light energy absorbed by the photosynthetic pigments, Fluorescence , phosphorescence,

Structural formula of the biliproteins that are present in the phycobilisomes, phycocyanin (black), and phycoerythrin (structural differences from phycocyanin are shown

Figure 2.15 Structural formula of the biliproteins that are present in the phycobilisomes, phycocyanin (black), and phycoerythrin (structural differences from phycocyanin are shown in red). The corresponding chromophores, phycocyanobilin and phycoerythrobilin are covalently bound to proteins via thioether linkages between the SH group of a cysteine residue of the protein and the vinyl group of the chromophore. The conjugated double bonds (red) show molecules with pigment-like character.

Figure 2.16 Absorption spectra of the phycobiliproteins phycoerythrin, phycocyanin, and allophycocyanin. For the sake of comparison

chlorophyll-a is shown

	Photosynthesis Playlist

1	Light-Harvesting: Photosynthetic Pigments and Exciton Transfer
2	Light-Harvesting: The Antenna Complex
3	Photosystem II Function: The P680 Reaction Center
4	The P680/P700 Special Pair Chlorophylls
5	Water Splitting: The Oxygen-Evolving Complex
6	Photosystem II and the Cytochrome b6f Complex Photosynthesis (Part 4)
7	Cytochrome b6f: Proton Pumping and ATP Synthesis
8	Photosystem I Functions: Electron Flow and Function
9	Plastocyanin, Ferredoxin, and Ferredoxin NADP+ Oxidoreductase (Photosynthesis Part 5)
10	Ferredoxin:NADP+ Oxidoreductase (Photosynthesis Part 6)
11	Ferredoxin:NADP+ Reductase: Generation of NADPH
12	Calvin Cycle: Carbon Fixation, Rubisco, and Rubisco Activase
13	Ethylene: Function and Synthesis in Plants
14	The Chlorophyll Cycle
15	Chlorophyll Catabolism and Pheophytin Synthesis
16	Rubisco in C3 Plants, 2-Phosphoglycolate, and Photorespiration
17	C4 Plant Metabolism: Spatial Separation
18	CAM Plant Metabolism: Temporal Separation

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substrate nutrition

video-Visualizing Soil Properties: Water Infiltration

OSU Extension – Containers and Media for the Nursery

Growing Media for Container Production in a Greenhouse or Nursery

Media Chemical Properties

Table 1. Typical ranges based on a saturated media extract (SME) analysis.
Parameter	Typical Range
pH	5.0-6.5
EC	. .500-3,000 umho/cm
NO3–N	. .40- 00 ppm
P	. .5-50 ppm
K	. .50- 00 ppm
Ca	. .30-1 0 ppm
Mg . . . . . . . . . . . . . . .	0-75 ppm
Na	. .4-80 ppm

Media pH is a critical issue because it plays a major role in determining the availability of many nutrients A common problem occurs in organic-based mixes when the pH falls below 5 0 Below this pH, the availability of Zn and Mn increases dramatically and often results in foliar toxicities from these elements While aluminum toxicity is recognized as a common concern in mineral soils at a low pH, this is usually not a problem in organic-based mixes

Cation Exchange Capacity (CEC)

CEC can vary widely depending on the type of component For example, perlite (1 5 meq/100 gm) and sand have very low CEC values relative to peat and vermiculite (125 meq/100 gm) components Some growers have started to use small volumes (2 to 15 percent) of clay-type amendments (e g , zeolites) in their soilless media These clay amendments may increase a medium’s nutrient retention and improve its physical properties It is important to understand which components have a higher CEC to help develop fertility programs and troubleshoot certain nutrient disorders

Soluble Salts

Similar to media pH, the level of soluble salts that may be tolerated is crop specific The extent of injury will be determined by the plant type, stage in production, longevity of exposure, concentration of salt and irriga- tion practices In general terms, fresh media without fertilizers should have a pour-through EC of less than 750 umho/cm The addition of fertilizer may mean this value is much higher and normal for a particular crop at a specific stage of production

Adjusting Media pH

Raising Media pH

In most cases, nursery and greenhouse growers need to be concerned about raising the media pH since most of the organic media components are acidic The most commonly used material is either calcitic (CaCO₃) or dolomitic limestone (mixture of CaCO₃ and MgCO₃) The amount of lime required will depend on the starting pH, the desired pH, the particle size of the limestone (i e , small particles faster acting than large ones), the type of media and the alkalinity of irrigation water used In general, lime rates generally fall in a range between 5 and

15 lbs/yd3 with rates below 8 pounds most common Calcitic or dolomitic limestone is most reactive when incorporated into the media prior to planting Note that many of the pelletized granular limestone materials are actually fine powders that have been glued together with a binder When these granules or prills are exposed to water, they fall apart into a fine powder These fine powders are faster acting than coarser prills but may also wash out of the bottom

of the pot if the media is coarse textured It is absolutely critical that growers know the starting pH of their mix and then monitor the pH over time to see how their fertilizer and irrigation water influence media pH

Other liming materials include calcium oxide (CaO; quick or burned lime, which is very reactive, caustic and more expensive), hydrated lime [Ca(OH)₂; also fast acting, caustic and more expensive], marl, egg or oyster shells and wood ash

Greenhouse growers may wish to try one of the following if the pH needs to be raised once the plants are in production The first option is to apply a flow- able limestone drench Start with a 1 quart per

100 gallons rate Avoid getting this mixture on the foliage if possible The second option involves injecting potassium bicarbonate into the irrigation water Continued use of this method may require a grower to switch to a lower potassium source fertilizer

If you are using liquid fertilization (fertigation), you can increase your media pH by switching from an acid-based fertilizer (high percentage of nitrogen in the ammoniacal form) to basic fertilizers that are based on a higher percentage of the nitrogen in the nitrate form

Lowering Media pH

Only rarely do growers using organic mixes express interest in lowering the media pH Generally, the problem develops from using an irrigation water source that has high alkalinity (>100 ppm CaCO₃) In those cases, growers typically choose to install an acid injector Others methods are selected if individual blocks of plants require a lower media pH

Materials such as elemental sulfur, ammonium sulfate and ferrous sulfate have all been used Caution must be used when considering using ammo- nium sulfate and ferrous sulfate as you need to account for the nitrogen and iron that accompanies these materials Aluminum sulfate is also an option but is used only for reducing pH around floristhydrangeas (Hydrangea macrophylla) Growers can also affect media pH by selecting specific forms of nitrogen The use of high ammoniacal-nitrogen based fertilizers can lower the media pH over time

Rate recommendations take into consideration the change in pH and type of media and may be obtained from grower manuals or your Cooperative Extension Service

Managing Substrate EC

Electrical conductivity (EC) is a good estimate for the total soluble salts in a media EC does not provide details on the type or amount of individual salts present High ECs can contribute to poor shoot and root growth The first objective is to determine the source for the elevated salts Typically, this will be from the irrigation water source or from the amount or type of fertilizer used Once the source has been identified, you will want to determine if you can reduce or eliminate that source Media salt concentra- tions are directly impacted by what is called the leaching fraction This value represents the percentage of water that leaves a container relative to what is applied High salt conditions can be effec- tively managed by keeping the leaching fraction high (20 to 30 percent) and not allowing pots to dry out The danger in keeping pots wet is that it can contribute to secondary problems with root rot organ- isms (Consult FSA 0 1, Irrigation Water for Greenhouses and Nurseries, for more information on irrigation water quality )

Disinfecting Media

Three methods are primarily considered for sterilization of media Sterilized media is common in plant propagation and greenhouse operations but is not usually considered in an outdoor nursery simply based on the volume of media required and the bene- fits derived Remember that certain amendments

(e g , perlite, vermiculite) are sterile and, therefore, do not require sterilization Composted pine bark and peat contain populations of suppressive microbes that might be eliminated by sterilization techniques

Steam Pasteurization

Steam pasteurization is commonly found in greenhouse or ground bed production The general recommendation is the exposure to steam (212°F) for 30 to 45 minutes The piles should be small enough so all sections reach at least 180°F Piles too big may take too long to achieve uniform heating You must have an appropriate thermometer handy to effectively monitor the temperature at various places in the pile or bed Over-steaming is possible and should be avoided since this kills beneficial organisms and may cause the release of toxic substances, especially when organic components are involved

An alternative pasteurization process, aerated steam, involves blowing a mixture of steam and air through the media Aerated steam (140° to 175°F) uses less energy and fewer beneficial organisms are likely to be harmed Steam pasteurization SHOULD NOT be used on media that has had slow-release fertilizer blended into it!

Chemical Fumigation

Chemical fumigation is usually limited to ground beds in cut flower production The primary chemicals used were methyl bromide and vapam Methyl bromide uses were phased out in 2005 Consult your Cooperative Extension Service for current recommendations

Solarization

Solarization is rarely used because the process may take up to one month even under summer condi- tions requiring tremendous planning for future media needs Solarization is accomplished by spreading moist media to a depth of 6 to 10 inches on a clean surface The pile or row is then covered with clear plastic sheets with the edges sealed to the surface to prevent the loss of heat and moisture

Media Physical Properties

Media physical properties may be determined using simple laboratory methods Media samples can be analyzed by commercial laboratories, or you can make the physical measurements yourself using simple tools Procedures for determining physical properties of horticultural substrates are available at https://projects.ncsu.edu/project/ hortsublab/pdf/porometer_manual.pdf.

4 Ways to Calculate Porosity – wikiHow

Weight (Bulk Density)

Media weight is kind of a double-edged sword Ideally growers would like a heavy mix when containers are on the ground in an outdoor nursery to minimize blow-over, but during plant movement and shipping, a lightweight mix is desired Weight or bulk density is usually expressed as lbs/ft3 and reported on a dry basis For outdoor container nurseries, dry bulk density of media might range between 12 to 24 lbs/ft3 (wet bulk density of 70 and 90 lbs/ft3) A nursery media that uses a significant percentage of mineral soil will have a dry bulk density of 40 to 50 ft3 For a greenhouse media, the dry bulk density will be lower and in the range of 8 to 18 lbs/ft3 Air-Filled Porosity

When we fill a container with media, the total volume of space in that container is filled with two things: the solid media components and the spaces or voids between all of the solids (Figure 1) Ideally the total volume of empty pores should be in the range of 50 to 70 percent This is referred to as total porosity The remaining container volume would be filled with the solid growing media (i e , 30 to 50 percent) The total porosity of a media is further composed of two parts: air and water Both components are critical for good plant growth, but not enough of either can limit growth

Figure 1. The container is filled with the solid media components and the spaces or voids between all of the solids.

For one quart and larger containers the air-filled porosity (percentage of pores filled with air) typically ranges from 10 to 20 percent by volume For a 280- plug tray, an acceptable range in air-filled porosity may fall in the range of 3 to 6 percent by volume Obviously, the container volume influences the inter- pretation of the acceptable values The higher the percentage of air-filled porosity, the more frequently watering will be required Propagation media where aeration and drainage are critical may have an air- filled porosity in the range of 15 to 25 percent

lumetric Moisture Content (sometimes referred to as Water-Holding Capacity)

As described above, the bulk volume of a container will be filled with either solids or pores These pores or voids are then either filled with water or air A typical range in values for the volumetric moisture content (percentage of the pores filled with water after allowing for free drainage) will be between 45 and 65 percent by volume Volumetric moisture content (VMC) gives a grower some indica- tion of how wet or how dry a media will be Sphagnum peat moss that retains water quite well typically has a VMC of 60 percent, while coarse sand, which does not hold water, might have a VMC of

25 percent As was the case with air-filled porosity, the actual values for VMC need to be interpreted relative to the height of the growing container For example, an acceptable VMC for a 6-inch container might be 45 percent, while for a plug tray 68 percent would be a more typical value The effect of media height on the saturated zone is illustrated in

Substrate Containers contain the same media. Notice, saturated zone (textured area at bottom of each container) is the same regardless of the container height.

Figure . Containers contain the same media. Notice, saturated zone (textured area at bottom of each container) is the same regardless of the container height.

All of these physical parameters can be determined in-house with the aid of a scale or balance The measurements can also be determined by an outside commercial laboratory or with the help of the Cooperative Extension Service

Graph-Electrical conductivity for various soil substrates

substrates-FSA-6098 Download

Containers and Media for the Nursery

Porosity: Measuring Pore Space

Pore space is expressed in terms of porosity, which is the percentage of pore space volume for a given substrate volume. You can determine the porosity, air space, and water space in your substrate by following the five steps below.

Determine the volume of the pot used to grow plants. You can usually obtain this from the manufacturer.
Add a water-tight liner (such as a polyethylene wrap) to the outer bottom of the pot. Tape the liner to the pot to prevent any material from draining through the holes.
Pour the substrate to the top of the pot. Make sure to compact the substrate as you would normally do during production. At this point, the substrate volume is equal to the volume of pot.
Add water to the top of the substrate, and carefully monitor how much you add. You want to add water until you completely saturate the substrate — that is, when you start to see a shiny layer of water over the substrate. Cover the pot with a lid and keep it aside for an hour. After one hour, check if you need to add a little more water to saturate the substrate. The total volume of water that you added to saturate the substrate

is equivalent to the total pore space volume.

Use this equation to determine porosity:

Porosity (%) = 100 x Volume of water added to fill Pot with substrate / Volume of Pot

Next, place the pot in a water-tight container and make holes at the bottom of the liner you added in step one. Your goal is to drain the water and collect it in the container. Let the pot drain for 10 to 15 minutes so the water can completely drain out. Then, measure the drained volume of water. This volume is equivalent to the air space volume in the substrate.

Use this equation to determine air space:

Air space = 100 x Volume of water drained / Volume of Pot

The water that is left in the substrate after it drains is the water space (also called maximum water-

holding capacity or container capacity). Once you’ve determined air space, you can determine water space with this equation:

Water space (%) = 100 – Air space(%)

Table 1. The following table provides information about the fractional volume of pore space, air space, and water space in two soilless substrates:

Note AFP has different definitions- some have air + water + solid -100

Component	Peat + Bark + Perlite + Vermiculite (P-B-P-V)	Peat + Perlite (P-P)
Solids (%)	40.9	38.6
Porosity (%)	59.1	61.4
Water space (as % total pore space)	72	62
Air space (as % total pore space)	28	38

Note AFP has different definitions- some have air + water + solid -100

substrate-pores Download

porosity-Growing-Media-AFP-testing-2013-Thea-Pobjoy Download

Soil Electrical Conductivity

Soil electrical conductivity (EC) measures the ability of soil water to carry electrical current. Electrical conductivity is an electrolytic process that takes place principally through water-filled pores. Cations (Ca2+, Mg2+, K+, Na+, and NH +) and anions (SO 2-, Cl-, NO -, and HCO -) from have higher EC than sandy outwash or alluvial deposits. Saline (ECe ≥ 4 dS/m) and sodic (sodium absorption ratio ≥ 13) soils are characterized by high EC. Scientific literature reported a relationship between EC values measured with commercial sensors and depths to claypan, bedrock, and fragipan. Microtopographic depressions in agricultural fields typically are wetter and accumulate salts dissolved in soil water carry electrical charges and conduct the electrical current. Consequently, the concentration of ions determines the EC of soils. In agriculture, EC has been used principally as a measure of soil salinity (table 1); however, in non-saline soils, EC can be an estimate of other soil properties, such as soil moisture and soil depth. EC is expressed in deciSiemens per meter (dS/m).

Factors Affecting

Inherent – Factors influencing the electrical conductivity of soils include the amount and type of soluble salts in solution, porosity, soil texture (especially clay content and mineralogy), soil moisture, and soil temperature. High levels of precipitation can flush soluble salts out of the soil and reduce EC. Conversely, in arid soils (with low levels of precipitation), soluble salts are more likely to accumulate in soil profiles resulting in high EC. Electrical conductivity decreases sharply when the temperature of soil water is below the freezing point (EC decreases about 2.2% per degree centigrade due to increased viscosity of water and decreased mobility of ions). In general, EC increases as clay content increases. Soils with clay dominated by high cation-exchange capacity (CEC) clay minerals (e.g., smectite) have higher EC than those with clay dominated by low CEC clay minerals (e.g., kaolinite). Arid soils with high content of soluble salt and exchangeable sodium generally exhibit extremely high EC. In soils where the water table is high and saline, water will rise by capillarity and increase salt concentration and EC in the soil surface layers.

It is generally accepted that the higher the porosity (the higher the soil moisture content), the greater the ability of soil to conduct electrical currents; that is, other properties being similar, the wetter the soil the higher the EC. Soil parent materials contribute to EC variability. Granites have lower EC than marine shales and clayey lacustrine deposits organic matter and nutrients and therefore have higher EC than surrounding higher lying, better drained areas.

Dynamic – Mineral soils enriched in organic matter, or with chemical fertilizers (e.g., NH₄OH) have higher CEC than non-enriched soils, because OM improves soil water holding capacity, and synthetic fertilizers augment salt content. Continuous application of municipal wastes on soil can increase soil EC in some cases. Electrical

conductivity has been used to infer the relative concentration, extent, and movement of animal wastes in soils. Because of its sensitivity to soluble salts, EC is an effective measure for assessing the contamination of surface and ground water. Although EC does not provide a direct measurement of specific ions or compounds, it has been correlated with concentrations of potassium, sodium, chloride, sulfate, ammonia, and nitrate in soils. Poor water infiltration can lead to poor drainage, waterlogging, and increased EC.

Relationship to Soil Function

Soil EC does not directly affect plant growth but has been used as an indirect indicator of the amount of nutrients available for plant uptake and salinity levels. EC has been used as a surrogate measure of salt concentration, organic

Table 1. Classes of salinity and EC (1 dS/m = 1 mmhos/cm; adapted from NRCS Soil Survey Handbook)

EC (dS/m)	Salinity Class
0 < 2	Non-saline
2 < 4	Very slightly saline
4 < 8	Slightly saline
8 < 16	Moderately saline
≥ 16	Strongly saline

matter, cation-exchange capacity, soil texture, soil thickness, nutrients (e.g., nitrate), water-holding capacity, and drainage conditions. In site-specific management and high-intensity soil surveys, EC is used to partition units of management, differentiate soil types, and predict soil fertility and crop yields. For example, farmers can use EC maps to apply different management strategies (e.g., N fertilizers) to sections of a field that have different types of soil. In some management units, high EC has been associated with high levels of nitrate and other selected soil nutrients (P, K, Ca, Mg, Mn, Zn, and Cu).

Most microorganisms are sensitive to salt (high EC). Actinomycetes and fungi are less sensitive than bacteria, except for halophyte (salt-tolerant) bacteria. Microbial processes, including respiration and nitrification, decline as EC increases (table 2).

Problems with Poor Soil EC Levels

High EC can serve as an indication of salinity (EC > 4 dS/m) problems, which impede crop growth (inability to absorb water even when present) and microbial activity (tables 2 and 3). Soils with high EC resulting from a high concentration of sodium generally have poor structure and drainage, and sodium becomes toxic to plants.

Improving Soil EC

Effective irrigation practices, which wash soluble salts out of soil and beyond the rooting depth, can decrease EC. Excessive irrigation and waterlogging should be avoided since a rising water table may bring soluble salts into the root zone. In arid climates, plant residue and mulch help soils to remain wetter and thus allow seasonal precipitation and irrigation to be more effective in leaching salts from the surface. To avoid the adverse effects of high EC (salinity) in irrigation water, the leaching requirement must be calculated for each crop. Leaching requirement is the fraction of water needed to flush excessive salt below the root zone, that is, the amount of additional water required to maintain a target salinity level. Adding organic matter,

such as manure and compost, increases EC by adding cations and anions and improving the water-holding capacity. In some cases, a combination of irrigation and drainage is necessary to lower salt concentration and EC. An EC water (ECw) ≤ 0.75 dS/m is considered good for irrigation water. Beyond this value, leaching or a combination of leaching and drainage will be necessary if the water is used.

Measuring Soil EC

The EC pocket meter is used to take measurements in the field. The method is described in the Soil Quality Test Kit Guide. Always calibrate the EC meter before use.

The pocket meter can be augmented by a probe that is placed directly into the soil to measure subsoil EC and NO – and make other estimates. NRCS soil scientists and agronomists use electromagnetic induction meters, not pocket EC meters, to map spatial variability of EC and associated soil properties at field scales. Special sensors are used for EC mapping for precision agriculture.

*Table 2. Influence of soil EC on microbial process in soils amended with NaCl or nitrate (adapted from Smith and Doran, 1996)*
Microbial process	Salt added	EC Range (dS/m)	Relative Decrease (%)	Threshold EC (1:1)
Respiration	NaCl	0.7 – 2.8	17 – 47	0.7
Decomposition	NaCl + alfalfa	0.7 – 2.9	2 – 25	0.7
Nitrification	soil + alfalfa	0.7 – 2.9	10 – 37	0.7
Denitrification	NO₃-N	1 – 1.8	32 – 88	1

*Table 3. Salt tolerance of crops and yield decrease beyond EC threshold (adapted from Smith and Doran, 1996)*
Crop species	Threshold EC 1:1 (dS/m)*	Percent yield decrease per unit EC beyond threshold EC
Alfalfa	1.1 – 1.4	7.3
Barley	4.5 – 5.7	5.0
Cotton	4.3 – 5.5	5.2
Peanut	1.4 – 1.8	29
Potato	1.0 – 1.2	12
Rice	1.7 – 2.1	12
Soybean	2.8 – 3.6	20
Tomato	1.4 – 1.8	9.9
Wheat	3.9 – 5.0	7.1
* Electrical conductivity of a 1:1 soil/water mixture relative to that of a saturated paste extract

Soil Quality Indicators: Soil Electrical Conductivity	USDA
Soil Electrical Conductivity	USDA
Using Soil Electrical Conductivity (EC) to Delineate Field Variation	Ohio State University
Pour-through Technique of Measuring Electrical Conductivity of the Substrate	Purdue University
Commercial Greenhouse Production: pH and Electrical Conductivity Measurements in Soilless Substrates	Purdue University

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A Nursery Friendly Method for Measuring Air Filled Porosity of Container Substrates

Bilderback-NC State

Porometer construction: Measuring air-filled porosity requires an apparatus called a porometer. Therefore, the first step is to construct porometers
Pre-moistening Substrate to Be Tested. Pre-moistening 12–24 h before testing is critical for achieving uniform and consistent results.
Packing Porometers with Substrate: After removing the plastic carton tops, individually weigh each porometer and record the weight. The weight of the plastic carton is subtracted from filled cartons as a “tare” weight to provide an accurate mass of substrate in each porometer. Next, overfill each porometer with potting substrate; tap each porometer firmly 3–5 times on a table or bench to eliminate air pockets and establish a bulk density
Saturate Substrate in Porometers: After packing, porometers are set upright in a vessel large enough for all of the test porometers to stand erect and tall enough to add water to the top of the porometers
Collecting and Measuring Drainage: Saturation of each porometer can be observed when water is seen at the surface of the substrate. Drainage from each porometer must be measured individually. This step may require practice. Fingers are used to prevent leaking from the drainage holes while the porometer is lifted from the saturation vessel and a pan is quickly placed under the drain holes. Porometers can be balanced on supports placed in the bottom of the drainage pan and allowed to fully drain. After draining has stopped, the drained volume is measured and recorded for each porometer
Calculating air filled porosity: The drainage volume is divided by the total volume for each porometer to determine a percent air-filled porosity (Table 1). Air-filled porosity measurements are added and divided by the number of porometers to obtain an average AFP for each test substrate. Changes in air filled porosity during a growing season or over a production cycle can be measured by placing porometers packed with substrate in containers which are set in nursery growing beds. Decomposition shrinkage should be measured and marked from the top of the porometer. The volume of the porometer marked at the surface of the substrate would be used as the new total volume and calculations followed as described above. If the important steps for pre- moistening samples and for packing to match the weight of each replicate sample in porometers are followed, consistent results can be accomplished.

Monitoring electrical conductivity in soils and growing media

What is EC? EC is a measure of the salinity (total salt level) of an aqueous solution. Pure, distilled water is a perfect insulator and it’s only because of dissolved ions that it can conduct electricity at all (Figure 1A). An EC meter measures the electrical charge carried by the ions that are dissolved in a solution— the more concentrated the ions, the higher the reading.

In nurseries, dissolved ions come from two sources (Figure 1B). First, all irrigation water contains some salt ions as rain water trickles through the soil and rocks.

The amount of the “background” salinity is a function of the local geology and climate. Soils and parent material have a major effect. Soils derived from marine sediments will contain high levels of sodium, chloride and sometimes boron. Water running through calcareous rocks or soils picks up calcium, magnesium and bicarbonate ions. Irrigation water from dry climates will have higher salinity than water from a humid climate. This only makes sense because, when water evaporates, the dissolved salts are left behind and the remaining solution would have a higher EC reading.

The second source of salinity in soils or growing media is from added fertilizers (Figure 1B). The release of salts varies considerably depending on how you are fertilizing. When fertigating, the soluble fertilizer that you inject into the irrigation water can be measured immediately. In fact, the best way to check the accuracy of your injector is to measure the EC of the applied fertigation solution. If you are incorporating controlled-release fertilizers into the soil or growing medium, however, then the salts are released according to fertilizer coating, water levels, and temperature. Most solid organic fertilizers release their nutrients very slowly and are less temperature or moisture dependent. Liquid organics release nutrients more rapidly but still much slower than soluble fertilizers.

EC Units. The physics and politics of this subject are complicated but think of it this way. We’re measuring electrical conductance which is the inverse of resistance. The unit of resistance is an ohm, and just to be cute, they call the unit of conductance a mho (pronounced “mow”), which is ohm spelled backwards. The most commonly used EC units in horticulture are micromhos per centimeter (µhos/cm), and the SI units of microsiemens per centimeter (µS/cm) which are equivalent. Because electron activity is strongly dependent on temperature, all EC measurement must be adjusted to a standard temperature of 77 °F (25 °C

Saturated Media Extract (SME). This technique is the laboratory standard that is used by commercial soil and water testing laboratories. If you are interested in absolute EC values, this is the only choice. The SME method uses saturation as the standard soil or media

The use of controlled-release fertilizers (CRF) has complicated the measurement of EC. Because the prills are very fragile, even collecting a sample or squeezing it can damage them. Broken prills will release all their fertilizer salts at once and artificially elevate the EC reading. Thus, some of the EC monitoring procedures should not be used when incorporating CRF

Pour-Through. This is a relatively new technique for measuring EC in containers, and works for all container types except for miniplugs where their short height stops the media solution from freely draining. It would also be impractical for very large containers which are difficult to move (Table 1). The pour-through process consists of 2 steps (Figure 4). First, medium in a container is progressively irrigated until saturated, and then left to stand for about 2 hours. Or, just do the procedure 2 hours after irrigation. Next, pour a volume of distilled water onto the media surface to produce about 100 ml of leachate. Of course, this depends on container volume and type of growing media. Make sure and apply the water slowly enough that it doesn’t run off and down the insides of the container. The idea is to have the applied water force out the solution surrounding the roots. The pour-through technique is ideal for growing media with controlled-release fertilizers because the prills are not squeezed or otherwise damaged (Table 1). Therefore, this method is ideal for outdoor growing compounds where controlled release fertilizers are the standard.

Figure 4 – The pour-through technique works for all containers except miniplugs and larges sizes, and is ideal when using controlled-release fertilizers

video-Pour Through Method for pH and EC

video-Hanna Lab – Set Up and Calibrate the Hanna Instruments pH, EC, TDS Combo Tester HI98129

video-Substrate Water Holding Capacity

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Formulae

Dry Bulk Density:

ρ_b = M_s / V_t

Where, ρ_b = Dry Bulk Density, M_s = Mass of Soil V_t = Volume

Wet Bulk Density:

P_t = (M_solid+M_liquid)/V_total

Where, P_t = Wet Bulk Density M_solid = Mass of Solids M_liquid = Mass of Liquid V_total = Total Volume

Moisture Content :

w = (M_w / M_s) * 100

Where, w = Moisture Content M_w = Mass of Water in Soil M_s = Dry Mass of Soil

Soil void ratio (e) is the ratio of the volume of voids to the volume of solids:

e = (V_v) / (V_s)

Where V_v is the volume of the voids (empty or filled with fluid), and V_s is the volume of solids.

soil porosity (n) is defined as the ratio of the volume of voids to the total volume of the soil. The posoity and the void ratio are inter-related as follows:

e = n /(1-n) , and n = e / (1+e)

The value of void ratio depends on the consistence and packing of the soil. It is directly affacted by compaction. Some typical values of void ratio for different soils are given below:

Description	USCS	Void ratio [-]	Reference
min	max	Specific value
Well graded gravel, sandy gravel, with little or no fines	GW	0.26	0.46		[1],
Poorly graded gravel, sandy gravel, with little or no fines	GP	0.26	0.46		[1],
Silty gravels, silty sandy gravels	GM	0.18	0.28		[1],
Gravel	(GW-GP)	0.30	0.60		[2],
Clayey gravels, clayey sandy gravels	GC	0.21	0.37		[1],
Glatial till, very mixed grained	(GC)	–	–	0.25	[4 cited in 5]
Well graded sands, gravelly sands, with little or no fines	SW	0.29	0.74		[1], [2],
Coarse sand	(SW)	0.35	0.75		[2],
Fine sand	(SW)	0.40	0.85		[2],
Poorly graded sands, gravelly sands, with little or no fines	SP	0.30	0.75		[1], [2],
Silty sands	SM	0.33	0.98		[1], [2],
Clayey sands	SC	0.17	0.59		[1],
Inorganic silts, silty or clayey fine sands, with slight plasticity	ML	0.26	1.28		[1],
Uniform inorganic silt	(ML)	0.40	1.10		[3],
Inorganic clays, silty clays, sandy clays of low plasticity	CL	0.41	0.69		[1],
Organic silts and organic silty clays of low plasticity	OL	0.74	2.26		[1], [3],
Silty or sandy clay	(CL-OL)	0.25	1.80		[3],
Inorganic silts of high plasticity	MH	1.14	2.10		[1],
Inorganic clays of high plasticity	CH	0.63	1.45		[1],
Soft glacial clay	–	–	–	1.20	[4 cited in 5]
Stiff glacial clay	–	–	–	0.60	[4 cited in 5]
Organic clays of high plasticity	OH	1.06	3.34		[1], [3],
Soft slightly organic clay	(OH-OL)	–	–	1.90	[4] cited in [5]
Peat and other highly organic soils	Pt	–	–		[4 cited in 5]
soft very organic clay	(Pt)	–	–	3.00	[4] cited in [5]

Factors Affecting Porosity of Soil:

Wide difference in the total pore space of various soils occurs depending upon the following several factors:

(i) Soil Structure:

A soil having granular and crumb structure contains more pore spaces than that of prismatic and platy soil structure. So well aggregated soil structure has greater pore space as compared to structure less or single grain soil.

(ii) Soil Texture:

In sandy soils the total pore space is small whereas in fine textured clay and clayey loam soils total pore space is high and there is a possibility of more granulation in clay soils.

(iii) Arrangement of Soil Particles:

When the sphere like particles is arrangement in columnar form (i.e. one after another on the surface forming column like shape) it gives the most open packing system resulting very low amount of pore spaces. When such particles are arranged in the pyramidal form it gives the most close packing system resulting high amount of pore spaces.

(iv) Organic Matter:

Soil containing high organic matter possesses high porosity because of well aggregate formation.

(v) Macro-Organisms:

Macro-organisms like earthworm, rodents, and insects etc. increase macro-pores in the soil.

(vi) Depth of Soil:

With the increase in depth of soil, the porosity will decrease because of compactness in the sub-soil.

(vii) Cropping:

Intensive crop cultivation tends to lower the porosity of soil as compared to fallow soils. The decrease in porosity may be due to reduction in organic matter content.

(viii) Puddling:

Due to puddling under sufficient soil moisture, the soil surface layer is made dense and compact. Eventually, the porosity of this surface soil is reduced by the infiltration of muddy surface materials.

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Although it may seem counter-intuitive, the small pore spaces of clay add up to more total void space than the fewer number of large pore spaces in sand. Consequently, in light rain or slow snowmelt, clay may be able to hold more water than sand.

However, water drains from clay soil more slowly than from sandy soils. So in successive rain events, clay soils may remain saturated between storms and therefore produce more runoff in the later rain events.

More References

Cuttings

cuttings-nursery-manual-native-plants.pdf

cuttings-PlantPropagationGraftingBuddin.pdf

cuttings-VegetativePropagationTechniques.pdf

CuttingsNutrient supply during cutting propagation.pdf

Hydroponic Nutrients

hydro-Hochmuth Hydroponic Tomato Fertilizer.pdf

hydro-EC-pH-management.pdf

hydro-Fertigation-Closed-Soilless-Culture.pdf

hydro-lettuce-cornell.pdf

hydro-low-pH-effects.pdf

hydro-nutrient-solutions.pdf

hydro-pH-floral-benchmark.pdf

hydro-pH-vegetables.pdf

hydro-tomato-oxygen.pdf

hydro-tomatoes-EC-pH.pdf

Nutrients

nutreints-vegetables-gh-mgt.pdf

nutrients-calcium-sprays-poinsettia-geranium.pdf

nutrients-deficiency-woody.pdf

nutrients-management-hydroponics.pdf

nutrients-Nitrogen-Physiology.pdf

Nutrients-recirculating solutions.pdf

nutrients-salinity-tolerance.pdf

nutrients-tissue-analysis-vege-crops.pdf

nutrients-tissues-levels-vegetable-crops-survey.pdf

planting substrates

substrates-containers.pdf

substrates-fertilizer-amendment.pdf

substrates-Leaching-pH.pdf

substrates-N-wood-chips.pdf

substrates-ph-chem-geranium.pdf

substrates-ph-effects-availability.pdf

substrates-tomato-EC-yield-quality.pdf

substrates-UArk-phys-chemm.pdf

substrates-wood-nutrients-requirements.pdf

soil reactions

Ion Exchange in Soil: Cation and Anion

Cation Exchange:

In a near neutral soil, calcium remains adsorbed on colloidal particle. Hydrogen ion (H⁺ ) generated as organic and mineral acids formed due to decomposition of organic matter. In colloid, hydrogen ion is adsorbed more strongly than is the calcium (Ca⁺⁺). The reaction takes place rapidly and the interchange of calcium and hydrogen is Chemically equivalent.

The reaction is as follows:

This phenomenon of the exchange of cations between soil and salt solution is known as Cation exchange or Base exchange and the cations that take part in this reaction are called exchangeable cations. Cation exchange reactions are reversible.

Hence, if some form of limestone or other basic calcium compound is applied to an acid soil, the reverse of the replacement just given above occurs. The active calcium ions replace the hydrogen and other cations by mass action. As a result, the clay becomes higher in exchangeable calcium and lower in adsorbed hydrogen and aluminium.

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Cation preferences

When the valence of the cations are equal (i.e. both +1 charge) the cation with the smallest hydrated radius is more strongly adsorbed. In the case of the monovalent cations of potassium and sodium, the potassium ion is more strongly adsorbed since it has a smaller hydrated radius and hence is more strongly adsorbed to the site of the negative charge. In comparison the sodium ion is so loosely held and so ready to hydrate that sodium rich soil will disperse.

Radius	Unit	Na⁺	K⁺	Mg²⁺	Ca²⁺	Al³⁺
Non-hydrated	nm	0.095	0.133	0.066	0.099	0.050
Hydrated	nm	0.360	0.330	0.430	0.410	0.480

This is similarly the case with the divalent cations of calcium and magnesium. Because the hydrated magnesium ion is larger than that of calcium, the magnesium ion is held more weakly and behaves in some instances in soil (i.e. when calcium is low) like sodium.

The charge of the cation and the size of the hydrated cation essentially govern the preferences of cation exchange equilibria. In summary, highly charged cations tend to be held more tightly than cations with less charge and secondly, cations with a small hydrated radius are bound more tightly and are less likely to be removed from the exchange complex. The combined influence of these two criteria can be summarized generally by the lysotrpoic series.

aluminium > calcium > magnesium > potassium, ammonium-NH⁴⁺ > sodium > hydrogen

It indicates, from left to right, the decreasing strength of adsorption of the various cations. As such, the less tightly held cations are located furthest from the surface of colloids and are most likely to be leached away or further down the profile most quickly. Conversely, the most strongly adsorbed cations will tend to move the slowest down through the profile.

The proportion and kinds of cations adsorbed on soil mineral particles and organic colloids is also a function of the concentration of cations in the soil solution. If the concentration of a cation in soil solution is high, there is an increased chance or tendency for that cation to be adsorbed.

This is the reason that dissolved gypsum (CaSO₄) is added to ameliorate sodic soil. In this case, the addition of dissolved gypsum increases the concentration of calcium in the soil solution and this leads to an increase in calcium ions on the exchange complex at the expense of exchangeable sodium.

The major source of cations in soil solution are from mineral weathering (i.e. primary minerals), mineralization of organic matter and addition of soil ameliorants (i.e. lime, gypsum, etc).

Soil Properties: Exchangeable Cations

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If a soil is treated with a liberal application of a fertilizer containing potassium chloride, following reaction may occur:

Some of the added potassium pushes its way into the colloidal complex and forces out equivalent quantities of calcium, hydrogen and other elements (e.g., M) which appear in the soil solution. The adsorption of the added potassium largely in an available condition. Hence, cation exchange is an important consideration for making already present nutrients in soils available to plants. Cation exchange also makes available the nutrients, applied in commercial fertilizers form.

Cation Exchange Capacity (C.E.C.):

The cation exchange capacity of a soil represents the capacity of the colloidal complex to exchange all its cations with the cations of the electrolyte solution (surrounding liquid). It also represents the total cation adsorbing capacity of a soil. Cation exchange in most soils increases with pH. At a very low pH value, C.E.C. is higher and at high pH, C.E.C. is relatively lower.

Factors affecting the Cation Exchange Capacity:

The following factors affect the cation exchange capacity:

(i) Soil texture:

Fine-textured (clay) soils tend to have higher cation exchange capacity (CEC) than sandy soils. Cation exchange capacity for clay soils usually exceeds 30 me/100 gm. while the value ranges from 0 to 5 for sandy soils.

(ii) Organic matter content:

Organic matter content of a soil affects the CEC. Higher organic matter content in a soil have higher CEC.

(iii) Amount and kind of clay:

Montmorillonite has higher CEC in comparison to illite or kaolinite clay.

(iv) pH:

The cation exchange capacity of most soils increases with pH. At very low pH value, the cation exchange capacity is also generally low. As the pH is raised, the negative charges on some 1 : 1 type silicate clay (Kaolinite), humus and Fe, Al oxides increases, thereby increasing the cation exchange capacity.

Milliequivalents:

The cation exchange capacity (C.E.C.) is expressed in terms of equivalents or more specifically, as milliequivalents per 100 grams. The term equivalent is defined as one gram atomic weight of hydrogen (or the amount of any other ion) that will combine with or displace this amount of hydrogen. For monovalent ions such as Na⁺, K⁺, NH₄⁺ and CI^–, the equivalent weight and atomic weight are the same since they can replace one H⁺ ion. Divalent cations such as Ca⁺⁺ and Mg⁺⁺ can take the place of two H⁺ions.

The milliequivalent weight of a substance is one thousandth of its atomic weight. Since the equivalent weight of hydrogen is about 1 gm., the term milliequivalent (meq) may be defined as 1 milligram of hydrogen. It indicates that other ions also may be expressed in terms of milliequivalents.

Consider calcium, for example. Ca has an atomic weight of 40 compared to 1 for hydrogen. Each Ca⁺⁺ ion has two charges and is thus, equivalent to two H⁺ ions. Therefore, the amount of calcium required to displace 1 mg of hydrogen is 40/2 = 20 mg (atomic wt. divided by 2 to obtain the equivalent wt.). This is the weight of 1 meq of calcium.

If 100 grams of a certain clay is capable of exchanging a total of 250 meq of calcium, the cation exchange capacity is 250/20 = 12.5 meq per 100 gm. The milliequivalent method of expression can be converted easily to practical field terms. For example, 1 meq of hydrogen can be replaced on the colloids by 1 meq of CaCO₃(limestone). The molecular weight of CaCO₃ is 100, it contains 2 equivalent weights (divalent).

Since the amount of CaCC₃ needed is only 1 meq wt., 100/2 = 50 mg will be needed to replace 1 mg of hydrogen (or 1 meq). In view of fact that 1 meq of H⁺ per 100 grams can be expressed as 20 pounds of hydrogen per grams of soil.

Expressed in the metric system this figure is 1100 kilograms per hectare. In general, the more clay there is in a soil, the higher the C.E.C. Sandy soils have, on the average 0.5 m.e. of C.E.C. per 100 gm. of soil, while for clay soils, it usually exceeds 30 m.e./100 gm.

Percentage Base Saturation of Soils:

Hydrogen and aluminium tend to dominate acid soils, both contributing to the concentration of H⁺ ions in the soil solution. Adsorbed hydrogen contributes directly to the H⁺ ion concentration in the soil. A⁺⁺⁺ions do so indirectly through hydrolysis.

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Base Saturation – Calculating Cation Exchange Capacity, Base Saturation, and Calcium Saturation

Base saturation is calculated as the percentage of CEC occupied by base cations. Figure 2 shows two soils with the same CEC, but the soil on the right has more base cations (in blue). Therefore, it has a higher base saturation. Base saturation is closely related to pH; as base saturation increases, pH increases.

Base Saturation (%) = (Base cations/CEC) x 100

Similarly, we can calculate the base saturation for each individual base cation. Calcium base saturation is calculated as the percentage of CEC occupied by calcium cations. In Figure 2, the soil on the right has twice as many calcium cations (Ca²⁺), thus a higher calcium saturation.

Calcium Saturation (%) = (Calcium cations/CEC) x 100

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Reactions are as follows:

Most of the other cations (Ca⁺⁺, Mg⁺⁺), called exchangeable bases, neutralize soil acidity. The proportion of the cation exchange capacity (C.E.C.) occupied by these bases is called the percentage base saturation. Thus, if the % base saturation is 80 in clay loam soil, 4/5th of the cation exchange capacity (20 meq) is satisfied by bases, the other by hydrogen and aluminium. Same as, 50% base saturation in clay soil having 20 meq C.E.C. x 1/2 (10 meq C.E.C.).of the C.E.C. is satisfied by bases likewise in sandy loam soil with a C.E.C. of only 10 meq, 80% base saturation satisfied, 4/5 of C.E.C.

A definite correlation exists between the percentage base saturation of a soil and its pH. As the base saturation is reduced as a result of loss of calcium in drainage, the pH is also lowered (more acidity)in a definite proportion.

Within the range pH 5 to 6, the ratio for humid temperate region mineral soils is roughly at 5% base saturation, change for every 0.10 change in pH. thus, if the percentage base saturation is 50% at pH 5.5, it should be 25% and 75% at pH 5.0 and 6.0.

Role of Cation Exchange:

Importance of exchangeable cations on plant nutrients is discussed below:

Cation exchange reactions are very important chemical reactions for the availability of plant nutrients in the soil. The capacity of soil to exchange cations is the best single index of soil fertility. Plant roots, when they come in contact with colloidal particles, absorb exchangeable cations directly by inter-exchange or contact exchange between the root hairs and colloidal complex.

(a) Nature and content of exchangeable bases:

The nature and content of exchangeable bases in a soil have an important bearing on its general properties. In all normal fertile soils the total exchangeable bases (Ca, Mg, K, Na) constitute about 80 to 90% of the cation adsorbing capacity. Exchangeable hydrogen is usually under 20%. In these soils, calcium forms the predominant exchangeable base, constituting 60 to 80% of the total exchangeable cation.

The predominance of exchangeable calcium give rise to Ca- clay which imparts a neutral reaction to the soil. The pH value varies from 6.5 to 7.5. When the proportion of exchangeable hydrogen (H) is high it gives rise to acid soil. In such soils, exchangeable calcium is correspondingly low, and in highly acid soils it is almost absent. In such cases the clay is saturated with hydrogen cations (H⁺) and forms H-clay. Acid soils are less fertile. It is called base unsaturated soil.

When exchangeable sodium form more than 10 to 15% of the total exchangeable cation it gives rise to alkaline soils. The pH value of such soils is usually greater than 8.0. When the proportion of exchangeable sodium exceeds these limits (or saturates the colloidal complex), the clay is turned into a Na-clay.

The soil is now highly alkaline and the pH value ranges from 9 to 12 Alkaline soils are also Jess fertile. Soils with a high calcium base saturation are in the most satisfactory physical and nutritional condition. A calcium dominated soil is granular in structure and ensure good drainage and aeration.

(b) Type of colloid:

Type of colloid affects the cation exchange. Montmorillonite colloid hold the calcium ion with greater tenacity than Kaolinite at a given base saturation. As a result, Kaolinite will liberate calcium much more readily than Montmorillonite.

(c) Associated ions:

Presence of exchangeable calcium in excessive quantities in a soil will limit the availability of potassium to plants. In same manner, high-exchangeable potassium may depress the availability of potassium to plants. In same manner, high- exchangeable potassium may depress the availability of magnesium.

(d) Adsorption of cations:

Colloidal clay (humus) hold in varying amount of plant nutrients (calcium, magnesium, potassium, nitrogen, phosphorus and most of the micronutrients) which are available to plant.

(e) Property of base exchange:

Base exchange (cation exchange) property checks leaching losses of available nutrients. On application of potassium sulphate fertilizer in the soil, potassium ions are held on the surface of colloids by cation exchange process. Subsequently, exchangeable potassium ions are directly available to plants.

Cation Exchange and Soil Fertility:

Cation exchange capacity is the best-index of soil fertility. By cation exchange, hydrogen ions from the root hairs and microorganisms replace nutrient cations from the exchange complex. The nutrient cations are forced into the soil solution where they can be assimilated by the adsorptive surface of roots and soil organisms, or they may be removed by drainage water.

(i) Cation saturation and Soil fertility:

Soil with a high calcium base saturation are the most satisfactory physical and nutritional condition. A calcium-dominated soil is granular in structure and porous. Calcium-dominated clay ensures good aeration and good drainage, thus increases fertility of the soils.

Base unsaturated soils are acidic in nature due to exchangeable hydrogen. These soils are less fertile. Base saturated soils with dominant sodium cations are alkaline in nature. Alkaline soils are not fertile due to de-flocculation, stickiness, hard to work, poor drainage and poor aeration.

(ii) Cation exchange and Soil fertility:

Due to the property of cation exchange (base exchange) the soluble inorganic fertilizer nutrients are not washed away from the soil. For example, ammonium sulphate fertilizer is added to the soil, ammonium ions are held on the surface of colloids by cation exchange. Ammonium ions are taken up by plants. This process checks nutrient losses by leaching and make the soil fertile. The cations Ca, Mg, K, and NH₄ are held on the colloidal surfaces and are readily available to plants.

(iii) Influence of complementary adsorbed cations and soil fertility:

The order of strength of adsorption, when the ions are present in equivalent quantities, is as follow:

Al³⁺> Ca²⁺> Mg²⁺> K⁺ = NH₄⁺> Na⁺

Consequently, a nutrient cation such as K⁺ is less tightly held by the colloids if the complementary ions are Al³⁺and H⁺ (acid soils) than if they are Mg⁺⁺ +Na⁺ (neutral to alkaline soils). The loosely held K⁺ ions are more readily available for absorption by plants or for leaching in acid soils.

There are also some nutrient “antagonisms”, which in certain soil cause inhibition of uptake of some cations by plants. Thus, potassium uptake by plants is limited by high levels of calcium in some soils. Likewise, high potassium levels are known to limit the uptake of magnesium even when significant quantities of magnesium are present in the soil.

Anion Exchange:

The process of anion exchange is similar to that of cation exchange. Under certain conditions hydrous oxides of iron and aluminium show evidence of having positive charges on their crystal surfaces. The positive charge of colloids are due to addition of hydrogen (H⁺) in hydroxyl group (OH^–) resulted in net positive charge (OH₂⁺). This + charge will attract anions (â€”).

The capacity for holding anions increases with the increase in acidity. The lower the pH the greater is the adsorption. All anions are not adsorbed equally readily. Some anions such as H₂ PO₄– are adsorbed very readily (quickly) at all pH values in the acid as well as alkaline range. Cl^– and SO₄– ions are adsorbed slightly at low pH but none at neutral soil, while NO₃– ions are not adsorbed at all. Hence, at the pH commonly prevailing in cultivated soilsâ€”nitrate (NO₃), chloride (Cl) and sulphate (SO₄) ions are easily lost by leaching.

In general, the relative order of anion exchange is:

OH > H₂PO_4–>SO_4–>NO_3–

Importance of Anion Exchange:

The phenomenon of anion exchange assumes importance in relation to phosphate ions and their fixation. The exchange is brought about mainly by the replacement of OH ions of the clay mineral.

The reaction is very similar to cation exchange:

The adsorption of phosphate ions by clay particles from soil solution reduces its availability to plants. This is known as phosphate-fixation. As the reaction is reversible, the phosphate ions again become available when they are replaced by OH ions released by substances like lime applied to soil to correct soil acidity.

Hence, the fixation is temporary. The whole of the phosphate adsorbed by clay is, however, not exchangeable, as even at pH, 7.0 and above. So, substantial quantities of phosphate ions are still retained by clay particles.

The OH ions originate not only from silicate clay minerals but also from hydrous oxides of iron and aluminium present in the soil. The phosphate ions, therefore, react with the hydrous oxides also and get fixed as in the case of silicate clay, forming insoluble hydroxy-phosphates of iron and aluminium.

If this reaction takes place under conditions of slight acidity it is reversible, and soluble phosphate is again liberated when hydroxy-phosphate comes in contact with ions. If the reaction takes place at a low pH under strongly acid conditions, the phosphate (ions) are irreversibly fixed and the totally unavailable for the use of plants.

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Chemistry and Behaviour of Phosphorus Present in Soil

Chemistry of Phosphorus:

1. Sorption Reactions:

The surfaces on which phosphate ions enter into sorption reactions of two types-surfaces of constant charge e.g. crystalline clay minerals and surfaces of variable charge including Fe³⁺ and Alâ€”oxides and organic matter where H⁺ and OH^– ions determine the surface charge and calcite (CaCO₃) in which Ca²⁺ and CO ions involve the charge development.

Besides, some other clay minerals including amorphous such as allophane also involves in the phosphate sorption.

Hydrated Fe and Al oxides are the most important surfaces of variable charge in most soils excepting peats and highly calcareous soils. These oxides have surfaces of negatively charged OH groups which take up and dissociate protons (H⁺) and hence they are amphoteric having either negative, zero or positive charge depending on pH.

The pH at which there are equal numbers of positive and negative charges on the surface is known as point of zero charge (PZC). At pH levels below the PZC, phosphorus and other anions like SO₄^2- and H3SiO₄^– are attracted to the positively charged oxide surfaces.

2. Precipitation Reactions:

Precipitation reactions mainly govern by the solubility product principles which are controlled by the pH of the system.

When some common phosphatic fertilizers like super phosphate, mono ammonium phosphate, Di-ammonium phosphate, some poly phosphates etc. are applied to the soil, within a very short time the released soluble phosphorus converts into very less soluble forms rendering unavailable and with time passes the strong insoluble phosphate fertilizer reaction products will form depending on the nature and type of soil as well as soil reaction.

In acid soils mono-calcium phosphate produces a number of substances like di-calcium phosphate (dihydrate and anhydrate), CaFe₂ (HPO₄)₄. 8H₂O; CaAl H(PO₄)₂.6H₂O etc. whereas in calcareous soils, di-calcium phosphate (CaHPO₄) is the dominant initial reaction product and in presence of excess amounts of calcium carbonate (CaCO₃), octacalcium phosphate may also form.

Further, when di-ammonium phosphate is applied to soils, the following reaction products viz. Ca₄ (PO₄)₃.3H₂O; Ca₂ (NH₄)₂ (NPO₄)₂.2H₂O, CaHPO₄-2H₂O; CaNH₄PO₄.H₂O; CaxH₂ (PO₄)6-5H₂O etc. will form. Dicalcium phosphate dihydrate is one of the most dominant reaction products formed in high-calcium soils followed by octacalcium phosphate.

When polyphosphate fertilizers are applied to soils it undergoes precipitation and adsorption reactions. In addition the orthophosphate present initially plus which formed by the hydrolysis of polyphosphates react with the soil components similar to that happened in orthophosphate compounds.

Hydrolysis of polyphosphates results in a stepwise breakdown forming orthophosphates and different short chain polyphosphate fragments. Then such short chain polyphosphates undergo further hydrolysis. However, reactions of polyphosphates in soil and the nature of substances produced are dependent upon the rate of their reversion back to orthophosphates.

Slow rate of hydrolysis permits condensed phosphates to sequester or form soluble complexes with soil cations and hence reduce phosphate retention in soils. Two mechanisms namely chemical and biological are involved in the hydrolysis of polyphosphates. In soils, where both mechanisms can function, the rate of hydrolysis will be rapid.

Enzymatic activity is the most important factor which controls the rate of hydrolysis. Phosphatases associated with plant roots and rhizosphere organisms are believed to be responsible for biological hydrolysis of pyro-and polyphosphates. Various factors like, temperature, soil pH, moisture, organic carbon content etc. can affect the transformation of polyphosphates.

Behaviour of Phosphorus:

Both organic and inorganic forms of phosphorus undergo transformation in soils leading to either release or retention of phosphorus. It is evident that decomposition of organic phosphorus substances gives both active and inactive substances.

The active substances are primarily the portions of the residues that have not yet been transformed into microbial products, whereas the inactive forms of phosphorus behave similarly to the resistant forms of nitrogen in humic acid.

1. Organic Phosphorus:

During mineralisation of organic phosphorus substances, the release of inorganic phosphorus takes place in the soil solution and such released phosphorus reacts very quickly with various soil components forming insoluble complex phosphatic compounds and there by unavailable to the plants.

Mineralisation of organic phosphorus is of three types:

(i) Based on the lowering of organic phosphorus level in soils due to long term cultivation.

(ii) Based on the results of short laboratory investigations decreasing the level of organic phosphorus with simultaneous increase in the amount of inorganic phosphorus in the soil and

(iii) Based on monitoring levels of soil organic phosphorus in the presence and absence of plants considering seasonal variation.

Mineralisation of organic phosphorus is carried by phosphatase enzymes and these enzymes are broad group of enzymes which catalyze the hydrolysis of both esters and anhydrides of phosphoric acid. However, there are a wide range of micro-organisms that are capable of mineralising (dephosphorylating) organic phosphorus on soils through their phosphatases activities.

Phosphatase activity of a soil is due to the combined functioning of the soil micro-organisms and any free enzymes present. Mineralisation of organic phosphorus is not entirely similar to that of organic carbon and nitrogen mineralisation and the mineralisation of organic phosphorus increases with an increase in soil pH but organic carbon and nitrogen mineralisation did not.

Most of the organic soil phosphates are present as inositol phosphate esters and these are prone to adsorption resulting less available in soils having higher adsorption capacity. The ultimate process by which organic phosphates are rendered available is by cleavage of inorganic phosphate by means of a phosphatase reaction.

The principle of this reaction is hydrolysis which is shown below:

For carrying out the mineralisation of organic phosphatic substances in soils it is essential to have some idea about C: N: P ratios in the soil. A carbon: nitrogen: phosphorus (C: N: P) ratio of 100: 10: 1 for soil organic matter has been advocated, but its values ranges from 229: 10: 0.39 to 71: 10: 3.05â€”depending on nature and type of soils.

C: P inorganic ratio â€“ Process Operates

200: 1 or less â€“ Mineralisation

Above 200: 1 but â€“ Neither net mineralisation nor

Less than 300: 1 â€“ Net immobilisation

300: 1 and above Immobilisation

A concentration of about 0.2% phosphorus is critical in the mineralisation of organic phosphorus substances. If the system contains less than this, net immobilisation takes place, as both the plant and the native soil phosphorus are utilised by micro-organisms. The transformation of P takes place both in upland (aerobic) and low land submerged (anaerobic) soils.

2. Inorganic Phosphorus:

It is evident that most of the soluble inorganic phosphorus either released from the mineralisation of organic phosphorus or applied as soluble phosphatic fertilizers are rendered unavailable to the plants and hardly 20% of the applied phosphatic fertilizers are available to the plant.

The reasons for such recovery are the conversions of soluble form of phosphorus to a form which is very less soluble through reactions with various soil components involving different mechanisms.

Such mechanism for the removal of phosphorus from the solution phase in the soil is known as “retention or fixation”. However, the retention of phosphorus in the soil involves various mechanisms namely, sorption and precipitation reactions.

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3 Main Forms of Potassium in Soils

Form # 1. Soil Solution Potassium:

It is recognised as the readily available form of potassium to the plants. The potassium availability in soils is controlled not only by the soil solution potassium but also by its buffering capacity (ability of a soil to maintain potassium intensity). The soil solution potassium (intensity, I) is maintained by the exchangeable potassium (quantity, Q) in a dynamic equilibrium.

The higher dQ/dl or the higher potassium buffer capacity indicates that during active period of crop growth, the potassium concentration in the soil solution will be depleted very rapidly. Soil solution potassium content usually higher in arid region and saline soil ranging from 3 to 156 ppm whereas the content of the same is lower in humid region soils ranging from 1 to 80 ppm.

Concentration of water soluble potassium may be as low as 8 ppm in deficient soils. However, under actual field conditions, the potassium concentration of soil solution varies with concentration and dilution processes brought about by evaporation and rainfall respectively.

The potentiality of soil solution K for plant growth and nutrition is influenced by the presence of other cations like Ca, Mg, and Al in acid soils and Na in salt affected soils. The activity ratio of potassium at equilibrium (AR_e^K) with respect to these above cations is a measure of the “intensity” of labile potassium in the soil indicating instantly available to plant roots.

Soils having same AR_e^K values may have different capacity in maintaining AR_e^K during depletion of K by crop uptake or leaching and hence for the K status of soils it is necessary to specify not only the status of potassium in the labile pool but also the way in which the intensity depends on the amount (quantity) of labile potassium present.

However, the detail discussion about the Q/I relationship of K is presented in the following section.

Buffer capacity indicates how intensity varies with quantity. A simple relationship between K intensity and K quantity for two soils (Soil X and Soil Y) having differential K adsorbing capacity is being depicted in Fig. 21.9. From the figure it is found that for both soils increasing intensity is accompanied by an increase in quantity.

Soil X, however, shows a steeper rise in the slope than that of soil Y. Where an equal amount of K is removed from both soils by plants a similar decrease in the quantity of (âˆ†Q) takes place. The consequent decrease in intensity (âˆ†l), however, varies greatly for both soils (âˆ†Ix and âˆ†I_Y).

This example shows that the two soils differ in their capacity of replenishing the soil solution with K. Soil X is better able to maintain the K concentration in the soil solution. Soil X is, therefore, more buffered than soil Y.

In quantitative terms the buffer capacity is expressed as the ratio âˆ†Q/âˆ†I as follows:

B_K = âˆ†Q/âˆ†I,

where B_K = buffer capacity of K in soils

The higher the ratio of âˆ†Q/âˆ†I, the more the soil is buffered. Usually, the rate of K uptake by plant roots is higher than the diffusive flux of K towards the roots. The K concentration at the root surface may decrease during the period of plant uptake.

Such decrease in K concentration is dependent on the K buffering capacity of the soil. If the buffer capacity is high, the decrease may be low because of efficient K replenishment of the soil solution.

Again, for spoils having poor K buffer capacity, the concentration of K at the root surface may decrease appreciably throughout the plant growth period. For optimum growth of the plant, the concentration of nutrients in soil solution should be maintained above a certain level.

This concentration is termed as the critical nutrient concentration (CNC) below which the yield of crop is decreased. The critical level of K in the bulk soil solution is related to the buffer capacity of K (buffer power). The critical concentration is higher, the lower K buffer capacity.

In addition to AR_e^K in assessing soil solution K, electro-ultra filtration (EUF) technique, a process of combination of electro dialysis and ultrafiltration, is used most satisfactorily for the characterisation of soil solution K (intensity) particularly in upland soils. The principle of EUF technique consists of utilizing the acceleration imposed upon ions by an electrical field for the separation of ions from soil colloids.

By adequate variation of voltage (50, 200 or 400 V) and timing (0-35 minutes), the total extractable K or any other nutrients can be separated into their water soluble and exchangeable forms with varying bonding energies.

Extraction and fractionation is done automatically. For the EUF-K fraction I (potassium in the extract obtained after 10 min of EUFâ€”the first 5 min at 50 V and the next 5 min at 200V) and total EUF-K fractions (sum of all potassium fractions obtained after 35 min of EUFâ€” the first 5 min at 50 V, the next 25 min at 200 V, and the last 5 min at 400 V).

The EUF-K fraction I considered as soil solution K (intensity factor) whiles the total EUF-K fractions as total amount of effectively available K (quantity factor). This electro- ultra filtration technique is better suited than that of AR_e^K in distinguishing soils of varying K availabilities.

The amount of K in the soil solution is very low to meet the demand of the crop throughout the growing period and therefore it is necessary for satisfactory potassium nutrition of crops the soil solution K must be continuously replenished from the exchangeable, non-exchangeable and mineral forms of K.

Form # 2. Exchangeable Potassium:

Potassium ion (K⁺) is held by soil colloids through electrostatic attraction similar to other cations. However, potassium held by soil colloids is easily displaced or exchanged when extracting the soil with neutral salt solutions. The amount of K exchanged varies with cations and usually neutral normal ammonium acetate solution is used for the purpose.

A small amount of potassium in this fraction occurs in soils (<1.0% of the total potassium). The distribution of potassium on soil colloids as well as soil solution depends upon nature and amounts of complementary cations, anion concentration and nature and characteristics of clay minerals.

As for an example, if a soil colloid is saturated with potassium and in that condition a neutral salt like calcium sulphate is applied then the following exchange reaction takes place:

Besides, if a soil is saturated with Al and Ca and in that conditions the application of muriate of potash gives the following exchange reaction:

When muriate of potash is applied to soils containing adsorbed calcium and aluminium, calcium is more easily replaced than aluminium by potassium. Coarse textured sandy soils having a greater base saturation lose very little of their exchangeable potassium by leaching as compared to soils containing low basic cations.

Liming is considered as the most common method of increasing the base saturation of soils which results the decrease in the loss of exchangeable potassium.

Sites for K Exchange:

It is evident that the exchangeable potassium on soil colloids is not homogeneous. Usually potassium is held at three binding sites of soil colloids namely p-(planar) position (outer surface of colloids, non-specific), e-(edge) position and i-(inner or inter layer) position (specific for K).

The amount of K held on p-position is in equilibrium with the soil solution K, while the amount of soil solution K in equilibrium with K held on e and i positions of soil colloids is low. However, under actual field situations, potassium concentrations in the soil pollution are probably the net result of three possible equilibria.

It is evident that the exchangeable form of potassium plays an important role in replenishing soil solution potassium removed by either intensive cropping or leaching losses.

In view of the above fact, it is very much essential to establish the quantity relationship between exchangeable K (Q quantity) and the activity of potassium in the soil solution (I intensity) in order to assess the availability of more labile potassium in soils to plants (Fig. 21.10).

The Q/I concept has been developed by Beckett which is used for predicting the status of potassium in soils. Different parameters of the above curve have some practical implications in relation to potassium in soils and plants.

âˆ†K = Amount through which the soil gains or loses potassium in bringing equilibrium (Q, quantity factor).

AR^K = Activity ratio of potassium (I, intensity factor).

AR_e^K = Activity ratio of potassium at equilibrium

âˆ†K_ex = Exchangeable or labile pool of potassium

K_SP = Specific sites for potassium

PBC^K = Potential buffering capacity

AR^K (Intensity Factor, I):

It is calculated from the determined concentration of calcium, magnesium, potassium and sodium correcting to the appropriate activities with the help of extended Debye-Huckel theory.

AR_e^K (Activity Ratio of K at Equilibrium):

It is a measure of availability or intensity of labile pool of potassium in soil and can be modified by potassium fertilization, being increased due to application of K fertilizers. However, the availability of potassium in soils can either be increased or decreased due to liming which modifies the AR_e^K values either favorably or adversely.

âˆ†K_ex:

It is used more successfully for the estimation of labile soil potassium held in plannar (p) positions. Greater values of labile potassium i.e. more negative (-K_ex) indicate a higher potassium release into the soil solution which results greater amount of potassium in the labile pool. However

the application of potassic fertilizers and lime in the cropped field have been found to be increased the amount of potassium in the labile pool.

K_sp (Specific Sites for Potassium):

It is a curved portion of the Q/I relationship while the linear portion of the curve (Q/I) is attributed to non-specific sites for potassium. Specific sites having high affinity for potassium are believed to exist on edges of clay minerals (e-positions) and in interlayer or wedge zones of weathered micas (i-positions).

Whereas non-specific sites for potassium are associated with planar surfaces of clay minerals (p-positions). The i-position has the greatest specificity for K⁺ which largely account for K⁺fixation in soils.

PBC^K (Potential Buffering Capacity of Potassium):

It is a measure of ability of a soil to maintain potassium concentration in the soil solution. The potential buffering capacity for potassium is proportional to the cation exchange capacity (CEC) of the soil that means, with an increase in CEC the value of PBC^K increases and vice-versa resulting from changes in AR^K values.

A high PBC^K value indicates a good potassium supplying power of soils whereas a low PBC^K value signifies very low potassium supplying power of soils indicating frequent potassium fertilization. However, the higher PBC^K value may be obtained due to time application which probably as a result of increase in pH-dependent cation-exchange capacity.

If PBC^K is low, small changes in exchangeable potassium produce large differences of potassium content in the soil solution. This value is very small coarse textured sandy soils where mainly organic matter is contributed to the CEC value. In such soils, extensive leaching, rapid plant growth etc. deplete available potassium within a few days.

In general, the relation between exchangeable and soil solution potassium is a good measure of the availability of the labile pool of potassium in soils to plants. The ability of a soil to maintain the activity ratio of K against depletion by crop uptake, leaching etc. is controlled by nature of the labile pool potassium as well as the rate of release of fixed potassium and diffusion and transport of K⁺ ions in the soil solution.

Form # 3. Non-Exchangeable and Mineral Form of Potassium:

Potassium in these forms is not readily available to the plants. However, non-exchangeable potassium pools not instantly available to plants, can contribute significantly to the maintenance of the labile pool of potassium in the soil. On the other hand, in some soils these fractions of potassium may become available as water-soluble and exchangeable forms are removed by leaching, crop uptake etc.

These forms of potassium are consisting of different K-bearing minerals namely primary minerals (K-feldspars) and micas (muscovites, biotites etc.), originating from the parent rock and secondary minerals (clays of the illitic group) formed by alteration of micas.

The main source of K⁺ for plants growing under natural conditions is from the weathering of K containing minerals mentioned above. In potash feldspars, potassium occurs in the interstices of the Si, Alâ€”O framework of the crystal lattice and held rigidly by covalent bonds. The weathering of feldspars starts at the surface of the particle.

Initially potassium is released by water and weak acids at a more rapid rate, However, with the progress of weathering, a Siâ€”Alâ€”O residue envelope is formed surrounding the un-weathered core. This layer reduces the rate of potassium loss from the mineral and hence protects K from further degradation.

Minerals of the mica type and also the secondary minerals of 2: 1 layer silicates vary in structure from feldspars and thereby these minerals also differ in their properties of releasing and binding potassium.

The micas consist of unit layers each containing two Si, Alâ€”O tetrahedral sheets between which is an M (Al, Fe, Mg)â€”O, OH octahedral sheet potassium (K⁺) ions occupy the approximately hexagonal spaces between the unit layers and as a result the distance between unit layers is relatively small i.e. 1.0 nm in micas.

The replacement of un-hydrated interlayer K⁺ by hydrated cations like Na⁺, Ca²⁺ or Mg²⁺ expands the mineral with an increase in the distance between the unit layers i.e., 1.4 nm in vermiculite (Fig. 21.11 and 21.12).

Usually K⁺ of the lattice is vulnerable to weathering and can diffuse out of the mineral in exchange for other cations. High H⁺ concentrations and low K⁺ concentrations in the soil favour the net release of non-exchangeable, inter layer K⁺.

This K⁺ release may be an exchange process associated with diffusion in which K⁺ adsorbed to i-positions of the inter layer zone is replaced by other large cations like Na⁺, Ca²⁺ and Mg²⁺ resulting an expansion of clay lattice and the formation of wedge zones (See in above figure).

“Frayed edge” or “wedge” zone formation is typical of weathering micas which results release of interlayer K⁺. The rate of release of K⁺ by weathering not only depends on the K content of the mineral, but also affected by structural variation between minerals.

The gradual release of potassium from positions of mica lattice results in the formation of illite (hydrous mica) and eventually vermiculite with accompanying gain of water or H₃O⁺and swelling of the lattice (Fig. 21.13).

There is also an increase in specific surface charge and CEC of clay minerals formed during the weathering of K containing minerals as well as transformation of mica. However, the applied soluble potassic fertilizers are converted to fixed or non- exchangeable forms of K and such conversions are affected by various factors. Besides fixation, the applied potassic fertilizers also undergo leaching loss from soils.

It is evident that a substantial amount of potassium can be lost through leaching in soils containing more amounts of sands due to flooding. However, in case of silty loam and clay loam soils, the loss of K through leaching is less because of fairly higher rate of adsorption of potassium by soil colloids. Again, in organic soils e.g. muck soils have high exchange capacities.

The bonding strength for cations like potassium is not great and the amount of exchangeable K tends to vary with the intensity of rainfall. Therefore, care should be taken for the supply of potassium to crops through its annual application.

Leaching losses can be reduced with the application of lime to the soil by maintaining a favourable pH level. Leaching losses of potassium frequently occurs in coarse textured sandy or organic soils particularly in areas of high rainfall.

plant fertilizer calculation videos 4

See Fertilizer Chemistry

Example Fertilizer Calculation N, P2o5, K20

bove is an example of a nutrient labels guaranteed analysis.
It tells us how much of each element is in the bag at percentage weight by volume (%w/v).
This provides us with enough information to establish a reasonably accurate ppm.

Note that analyzing the ppm from fertilizer labels won’t provide 100% accurate ppms.
Fertilizers sold worldwide are often only required to be listed accurately to within 0.4%.

Regulations around the world require that NPK.. values be presented somewhat ambiguously.
Therefore, listings for the same nutrient may appear to vary on a country-by-country basis.
For example, when looking at our labels guaranteed analysis you will find note that it states;

Available Phosphate (P2O5)……….15.0%
Available Potash (K2O)…………..14.0%

This information becomes important when interpreting the guaranteed analysis.

That is, it is important to note that the P and K numbers found on the guaranteed analysis do not always reflect the actual amounts of elemental phosphorous and potassium by %.

With our label, this is the case and P is listed as P2O5 (phosphorous pentoxide) and K is listed as K2O (potassium oxide) percentage.

When phosphorus is listed as P2O5 it is only 43% elemental P and when potassium is listed as K2O it is only 83% elemental K.

Therefore, when this system is in use, a 5-15-14 NPK ratio truly reflects elemental NPK 5-6.45-11.62.

N = 5
P = 15 * 0.43 = 6.45
K = 14 * 0.83 = 11.62

Additionally, other nutrients such as calcium (Ca), magnesium (Mg) and sulfur (S) can be listed in their oxide form (CaO, MgO, SO3) or in elemental form, or both.
To convert other nutrient listings that may appear on some labels use these equations.

CaO to Ca multiply by 0.714
MgO to Mg multiply by 0.6031
SO3 to S multiply by 0.4

Percentage Weight by Volume (%w/v)

A simple way of understanding how to convert a %w/v listing found on the guaranteed analysis into grams per litre is by understanding that 1ml of RO water weighs 1gram.

Percentage weight by volume %w/v refers to the total weight of elements contained within a finished concentrate of a given total volume.

For example, 5% of nitrogen added to 1 litre(1000ml) of RO water would mean that there is 50grams of N in the water.

1000 (ml) * 0.05 (5% nitrogen) = 50 (grams of N)

Converting %w/v to ppm and ppm to %w/v

To establish ppm from %w/v you simply need to multiply by 10000.
5% (nitrogen) * 10000 = 50000 (ppm)
To establish %w/v from ppm you simply need to divide by 10000.
50000 (ppm) / 10000 = 5% (nitrogen)

To establish the concentration of individual elements in the water, the guaranteed analysis (%w/v) should first be converted into ppm, then multiplied by the usage rate (per litre), then divided by 1000 (ml).

For example, if a nutrient lists 5% nitrogen, when it is used at 5grams per 4 litres it will yield 62.5 ppm of nitrogen per litre.

Step 1 : 5% (nitrogen) * 10000 = 50000 (ppm)
Step 2 : 5grams / 4L = 1.25g/litre
Step 3 : (50000 (ppm) * 1.25g/litre) / 1000ml (1 litre) = 62.5ppm of nitrogen per litre (1000ml)

Doing the math

Using what we’ve learned, we’re finally ready to find the ppm of our fertilizer.

5 * 10000 = 50k nitrogen ppm
6.45 * 10000 = 64.5k phosphorus ppm
11.62 * 10000 = 116.2k potassium
5 * 10000 = 50k calcium ppm
3.5 * 10000 = 35k magnesium ppm
4 * 10000 = 40k sulfur ppm
0.1 * 10000 = 1k iron ppm

50k + 64.5k + 116.2k + 50k + 35k + 40k + 1k = 356,7k or 356700 ppm.
(356700ppm * 1.25g/litre) / 1000ml = 445ppm(~0.9EC) per litre.

How Much Phosphorus and Potassium are Really in Your Fertilizer?

Fertilizer Calculations for Greenhouse Crops

Understanding phosphorus fertilizers

Common Fertilizers Table

Table 1: Percentages of water-soluble and available phosphate in several common fertilizer source

P₂O₅ source	N	Total	AvailableP₂O₅	Water soluble* P₂O₅
Superphosphate (OSP)	0%	21%	20%	85%
Concentrated Superphosphate (CSP)	0%	45%	45%	85%
Monoammonium Phosphate (MAP)	11%	49%	48%	82%
Diammonium Phosphate (DAP)	18%	47%	46%	90%
Ammonium Polyphosphate (APP)	10%	34%	34%	100%
Rock Phosphate	0%	34%	38%	0%
*Water-soluble data are a percent of the total P₂O₅.
Source: Ohio Cooperative Extension Service.

Other Nutrients

Potash

Common name	Chemical name (Formula)
Potash fertilizer	c.1942 potassium carbonate (K₂CO₃); c.1950 any one or more of potassium chloride (KCl), potassium sulfate (K₂SO₄) or potassium nitrate (KNO₃).^[9][10] Does not contain potassium oxide (K₂O), which plants do not take up.^[11] However, the amount of potassium is often reported as K₂O equivalent (that is, how much it would be if in K₂O form), to allow apples-to-apples comparison between different fertilizers using different types of potash.
Nitrate of potash or saltpeter	potassium nitrate (KNO₃)
Sulfate of potash (SOP)	potassium sulfate (K₂SO₄)
Permanganate of potash	potassium permanganate (KMnO₄)

Potassium oxide (K₂ O) is an ionic compound of potassium and oxygen. The chemical formula K₂O (or simply ‘K’) is used in several industrial contexts: the N-P-K numbers for fertilizers,

P2o5

Phosphorus pentoxide is a chemical compound with molecular formula P₄O₁₀ (with its common name derived from its empirical formula, P₂O₅).

The phosphate or orthophosphate ion [PO ₄]³⁻is derived from phosphoric acid by the removal of three protons H⁺

Lime

Agricultural lime, also called aglime, agricultural limestone, garden lime or liming, is a soil additive made from pulverized limestone or chalk. The primary active component is calcium carbonate.. Calcium oxide (CaO), is commonly known as quicklim.e

Calcareous (/kælˈkɛəriəs/) is an adjective meaning “mostly or partly composed of calcium carbonate“, in other words, containing lime or being chalky.

Calcium carbonate shares the typical properties of other carbonates. Notably it

reacts with acids, releasing carbon dioxide (technically speaking, carbonic acid, but that disintegrates quickly to CO₂ and H₂O):

CaCO₃(s) + 2 H⁺(aq) → Ca²⁺(aq) + CO₂(g) + H₂O(l)

Calcium carbonate reacts with water that is saturated with carbon dioxide to form the soluble calcium bicarbonate.

CaCO₃(s) + CO₂(g) + H₂O(l) → Ca(HCO₃)₂(aq)

Agriculture and aquaculture

Agricultural lime, powdered chalk or limestone, is used as a cheap method for neutralising acidic soil, making it suitable for planting, also used in aquaculture industry for pH regulation of pond soil before initiating culture.^[54]

Nitrogen

Ammonia is a compound of nitrogen and hydrogen with the formula NH₃.

The ammonium cation is a positively charged polyatomic ion with the chemical formula NH⁺₄. It is formed by the protonation of ammonia (NH₃).

Urea, also known as carbamide, is an organic compound with chemical formula CO(NH₂)₂.

soil reactions

plant nutrients

Nutrient Management pdfs

GroAlert-geranium-nutrition-problems.pdf

UF-IFAS-GTO1 Substrate pH.pdf

UF-IFAS-GTO3 Onsite Soil Testing.pdf

UF-nutrition-Chapter 1 Introduction Course Text-1.pdf

UF-nutrition-Chapter 2 Fertilizers Course Text-1.pdf

UF-nutrition-Chapter 2a Supplying Fertilizer Course Text-1.pdf

UF-nutrition-Chapter 3 Managing Nutrient Level Course Text-1.pdf

plant nutrients

nutrients

plant chemistry

PDF excerpts

Plant and Soil Science Texts
Introduction to Botany (Shipunov)	Botany.pdf
The Biology of Soil	SOIL-BIO-Of-Soil.pdf
The Chemistry of Soils	SOIL-CHEM.pdf
The Chemistry of Soils	SOIL-Science-simplif..>
Introduction to Environmental Soil Physics	SOIL-enviro-physics-..>
	SOIL-enviro-physics-..>
	SOIL-enviro-physics-..>
Fundamentals of Plant Pathology	plant-Pathology-fund..>
Plant Pathology	plant-pathology.pdf
Plant Physiology	plant-physio-01-04.pdf
	plant-physio-05-10.pdf
	plant-physio-11-15.pdf
	plant-physio-16-20.pdf
	plant-physio-21-25.pdf
Crop Physiology	plant-physio-crop.pdf
Plant Biochemistry	plant Biochemistry 4..>
Introductory Plant Biology	plant Biology.pdf

Fertilizer Chemistry

Agricultural Salts

	N	P2O5	K2O	Ca	Ng	S
Ammonium Nitrate – NH₄ NO₃.	34	0	0	0	0	0
Ammonium Phosphate – (NH₄)₃PO₄.or ADP-MAP (NH₄)(H₂PO₄).	12	61	0	0	0	0
Ammonium Sulfate – (NH₄)₂SO₄	21	0	0	0	0	24
Calcium Nitrate – Ca(NO₃)₂ or CaN₂O₆	15	0	0	19	0	0
Magnesium Nitrate – Mg(NO₃)₂(	11	0	0	0	9	0
Magnesium Sulfate – MgSO ₄,	0	0	0	0	10	13
Potassium Nitrate – K N O ₃.	13	0	46	0	0	0
Urea – CO(NH₂)₂. carbamide	46	0	0	0	0	0

Lewis Dot Structure for the Ammonium Ion

Nitrate Ion Lewis Structure: How to Draw the Lewis Structure for Nitrate Ion

How to Draw the Lewis Dot Structure for NH4NO3: Ammonium nitrate

How to Write the Formula for Ammonium nitrate

PO4 3- Lewis Structure: How to Draw the Lewis Structure for PO4 3-

How to Write the Formula for Ammonium phosphate

How to draw the (NH4)3PO4 Lewis Dot Structure (Ammonium Phosphate)

How to Draw the Lewis Structure for the Sulfate Ion

How to Write the Formula for Ammonium sulfate

How to draw the (NH4)2SO4 Lewis Dot Structure (Ammonium Sulfate)

How to Draw the Lewis Dot Structure for Mg(NO3)2 : Magnesium nitrate

How to Draw the Lewis Dot Structure for KNO3 (Potassium Nitrate)

How to Draw the Lewis Dot Structure for CH4N2O / CO(NH2)2 : Urea

================================

Notes

Salts- In chemistry, a salt is a chemical compound consisting of an ionic assembly of cations and anions.^[1] Salts are composed of related numbers of cations (positively charged ions) and anions (negatively charged ions) so that the product is electrically neutral (without a net charge). Often a salt is an ionic compound in which the cation is a metal and anion is a nonmetal or group of nonmetals.

An oxyacid, oxoacid, or ternary acid is an acid that contains oxygen. Specifically, it is a compound that contains hydrogen, oxygen, and at least one other element, with at least one hydrogen atom bonded to oxygen that can dissociate to produce the H⁺ cation and the anion of the acid.^[1]

Element group	Element (central atom)	Oxidation state	Acid formula	Acid name^[8][9]	Anion formula	Anion name
7	Manganese	+7	HMnO ₄	Permanganic acid	MnO⁻₄	Permanganate
+6	H ₂MnO ₄	Manganic acid	MnO²⁻ ₄	Manganate
8	Iron	+6	H₂FeO₄	Ferric acid	FeO₄^2–	Ferrate
13	Boron	+3	H ₃BO ₃	Boric acid (formerly orthoboric acid)^[10]	BO³⁻ ₃	Borate (formerly orthoborate)
14	Carbon	+4	H ₂CO ₃	Carbonic acid	CO²⁻₃	Carbonate
Silicon	+4	H ₄SiO ₄	Silicic acid (formerly orthosilicic acid)^[10]	SiO⁴⁻₄	Silicate (formerly orthosilicate)
14, 15	Carbon, nitrogen	+4, −3	HOCN	Cyanic acid	OCN⁻	Cyanate
15	Nitrogen	+5	HNO ₃	Nitric acid	NO⁻₃	Nitrate
+3	HNO ₂	Nitrous acid	NO⁻₂	Nitrite
Phosphorus	+5	H ₃PO ₄	Phosphoric acid (formerly orthophosphoric acid)^[10]	PO³⁻₄	Phosphate (orthophosphate)
H ₃PO ₅	Peroxomonophosphoric acid	PO³⁻₃	Peroxomonophosphate
+5, +3	(HO) ₂POPO(OH) ₂	Diphosphoric(III,V) acid	O ₂POPOO²⁻ ₂	Diphosphate(III,V)
16	Sulfur	+6	H ₂SO ₄	Sulfuric acid	SO²⁻ ₄	Sulfate
H ₂S ₂O ₇	Disulfuric acid	S ₂O²⁻ ₇	Disulfate
17	Chlorine	+7	HClO ₄	Perchloric acid	ClO⁻ ₄	Perchlorate
+5	HClO ₃	Chloric acid	ClO⁻ ₃	Chlorate

Hydroxyl Group Definition

A hydroxyl group is a functional group that attaches to some molecules containing an oxygen and hydrogen atom, bonded together. Also spelled hydroxy, this functional group provides important functions to both alcohols and carboxylic acids. Alcohols are chains of carbon molecules with a functional hydroxyl group side chain. The electronegativity of the oxygen adds a slight polarity to alcohols, which is why they are able to interact with other polar molecules such as water and some solutes. Below is a general alcohol which contains a hydroxyl group. The oxygen is the red atom, while the hydrogen is represented by the grey atom. The R represent any generic carbon chain.

Carboxylic acids contain a hydroxyl group within their functional carboxyl group. A carboxyl group consists of a carbonyl group bonded to a hydroxyl group. A carbonyl group is simply a carbon double bonded to an oxygen. These two functional groups together create an extremely reactive molecule, which is prone to forming new carbon-carbon bonds. Along with alcohols, carboxylic acids are commonly seen in nature. A generic carboxylic acid with its hydroxyl group can be seen below.

Besides these two large classes of molecules that are functionally dependent on the hydroxyl group, many other molecules contain hydroxyl groups. As mentioned, a large part of the action caused by the hydroxyl group is due to the electronegativity of the oxygen. Because oxygen has a stronger attraction with the electrons bonding hydrogen to the molecule, the hydroxyl group can easily lose the hydrogen to an atom that will share electrons more equally. When this happens, the oxygen takes on a much more negative electrical energy, and can donate the extra electrons it has to a number of reactions. Biological organisms use this property of oxygen to help connect and disconnect chains of carbon molecules, which hold energy the organism can use to power cellular functions.

Related Biology Terms

Carboxyl group – A carbon doubled bonded to an oxygen and also bonded to a hydroxyl group.

Carbonyl group – A carbon double bonded to an oxygen and any other molecules, including more carbons.

Electronegativity – The attraction that an atom has for electrons, compared to the other types of atoms that it shares electrons with in covalent bonds.

Polarity – The property of a molecule that arises from the stable differentiation of electrical poles across a molecule or part of a molecule.

Nitric Acid- HNO₃

Ammonium nitrate is the ammonium salt of nitric acid. It has a role as a fertilizer, an explosive and an oxidising agent. It is an inorganic molecular entity, an ammonium salt and an inorganic nitrate salt.

Phosphoric acid, H₃PO₄ or H₃O₄P , is a phosphorus oxoacid that consists of one oxo and three hydroxy groups joined covalently to a central phosphorus atom. It has a role as a solvent, a human metabolite, an algal metabolite and a fertilizer. It is a conjugate acid of a dihydrogenphosphate and a phosphate ion.

Ammonium dihydrogen phosphate is the ammonium salt of phosphoric acid (molar ratio 1:1). It has a role as a fertilizer. It contains a dihydrogenphosphate.

Sulfuric acid, H₂SO₄ or H₂O₄S , is a sulfur oxoacid that consists of two oxo and two hydroxy groups joined covalently to a central sulfur atom. It has a role as a catalyst. It is a conjugate acid of a hydrogensulfate

Ammonium sulfate, (NH₄)₂SO_4, is an inorganic sulfate salt obtained by reaction of sulfuric acid with two equivalents of ammonia. A high-melting (decomposes above 280℃) white solid which is very soluble in water

Calcium nitrate, is inorganic nitrate salt of calcium. It has a role as a fertilizer. It is an inorganic nitrate salt and a calcium salt. It contains a calcium(2+).

NO₃^– Nitrate is a nitrogen oxoanion formed by loss of a proton from nitric acid.

Magnesium sulfate is a magnesium salt having sulfate as the counterion.. It is a magnesium salt and a metal sulfate.

Potassium nitrate is the inorganic nitrate salt of potassium. It has a role as a fertilizer. It is a potassium salt and an inorganic nitrate salt.

Urea is a carbonyl group with two C-bound amine groups. It has a role as a flour treatment agent, a human metabolite, a Daphnia magna metabolite, a Saccharomyces cerevisiae metabolite, an Escherichia coli metabolite, a mouse metabolite and a fertilizer. It is a monocarboxylic acid amide and a one-carbon compound. It derives from a carbonic acid. It is a tautomer of a carbamimidic acid.

Al₂(SO₄)₃. Aluminum Sulfate Anhydrous is an aluminum salt

Agricultural lime, Ca CO₃. ,also called aglime, agricultural limestone, garden lime or liming, is a soil additive made from pulverized limestone or chalk. The primary active component is calcium carbonate.

—-

Calcareous Soils	Plant Nutrition
Cation and Anion Exchange Capacity	plant videos 1 chemistry
Chemistry pH	plant videos 2
Fertilizer Chemistry	plant videos 3 physiology
greenhouse tech	plant water and transport
Micronutrients	Soil and Roots
nutrient table	soil science
Plant Cell Biology	spring-lake
Plant Cell Physiology	Stylistics
plant nutrients

nutrient table

N P K, C M S, I M Z, C B M.

Nutrient	Concentration	function	Uptake form	Mobility in Plant	Mobility in Soil	Acid Deficiency	Alkaline Deficiency
gas and water			C, H, O Carbon, Hydrogen, Oxygen
Carbon ©	50% dry weight	present in all macromolecules	CO₂, H₂CO₃ Carbonic acid
Hydrogen (H)	h2o is >80% plant weight	part of many organic compounds and also forms water	Hydron H⁺, OH^–Hydroxide , H₂O
Oxygen (O)		necessary for cellular respiration; plants use oxygen to store energy in the form of ATP.	O₂ oxygen gas
MACRO N P K C M S , (Ca Mg S)			N, P, K Ca Mg S Nitrogen, Phosporous, Potassium ; Calcium, Magnesium, Sulfur
Nitrogen (N)	4.00%	A component of chlorophyll, nucleic acids, proteins, and enzymes	NO₃^–Nitrate^, NH₄⁺ammonium ion, also see Urea CH₄N₂O , and Ammonium nitrate (NH₄NO₃)		Mobile as NO₃^–, immobile as NH₄⁺	y	Volatilization Urea
Phosphorus	0.50%	Required to store and transport energy	HPO₄^2-Hydrogen phosphate, H₂PO₄^– Dihydrogen phosphate		Immobile	y	y
Potassium	4.00%	Acts as a osmotic regulator in water absorption and loss by the plant.	K⁺			y
Calcium	1.00%	Cell structure, secondary plant hormone	Ca²⁺, see also Lime calcium carbonate (CaCO₃).	Low
Magnesium	0.50%	Central ion in the chlorophyll molecule	Mg²⁺		Immobile	y
Sulfur	0.50%	A component of nucleic acids and proteins	SO₄^– sulfate ion			y
MICROI I M Z , ( Fe Mn Z) C B M , (Cu B Mo)			Fe, Mn, Zn ; Cu, B, Mo Iron, Manganese, Zinc, Copper, Boron, Molybdenum
Iron	200 ppm	Required for chlorophyll synthesis and energy transferring pathways	Fe²⁺, Fe³⁺	Low	Immobile		y
Manganese	200 ppm	Required for chlorophyll production and energy transferring pathways	Mn²⁺	Low			y
Zinc	30 ppm	Activates enzymes	Zn²⁺	Low	Immobile		y
Copper	10 ppm	Involved in respiration and oxidation/reduction reactions	Cu²⁺	Low	Immobile		y
Boron	60 ppm	cell division and differentiation of young tissue	H₃BO₃Boric Acid_, BO₃^–Borate	Low			y
Molybdenum	1 ppm	Involved in nitrogen metabolism	MoO₄^–Molybdate	Low		y
other S CS C N
Sodium	500 ppm	Osmotic regulator	Na⁺			10⁻³ g	mg milligram
Chlorine	0.10%	Required for photosynthesis	Cl^–			10³ g	kg kilogram
Silicon	0.05-0.15%	Pathogen defense, drought and heat tolerance	H4SiO4			10⁻⁶ g	µg microgram (mcg)
Cobalt			Co²⁺	Low		1 ppm = 1/1,000,000 =0.000001 = 0.0001% = 1 mg/kg
Nickel			Ni²⁺			1 ppm = 1/1,000,000 =0.000001 = 0.0001% = 1 mg/kg

Refrences –

Nutrient Uptake in Plants, Smart Fertilizer

Soil analysis: key to nutrient management planning, PDA

The most commonly found nutrient deficiency and toxicity symptoms are presented in the table below:

Nutrient	Deficiency Symptoms	Toxicity Symptoms
Nitrogen (N)	Stunted growth and restricted growth of lateral shoots. Plants express general chlorosis of the entire plant to light green and yellowing of older leaves which proceeds to younger leaves. Older leaves become necrotic and defoliate early	Plants are stunted, deep green in color, and secondary shoot development is poor. High N causes vegetative bud formation instead of reproductive bud formation. Ammonium toxicity can cause roots to turn brown, with necrotic root tips; reduce plant growth; necrotic lesions occur on stem and leaves; vascular browning occurs in stems and roots.
Phosphorus (P)	Stunted growth. Purplish coloration of older leaves in some plants. Dark green coloration with tips of leaves dying. Delayed maturity, Poor fruit and seed development.	Excess P in the plant can cause iron and zinc deficiencies.
Potassium (K)	Leaf margins turn chlorotic and then necrotic. Tip and marginal burn starting on mature leaves. Lower leaves turn yellow. Weak stalks and plant lodge easily. Slow growth.	High amounts of K can cause calcium (Ca), magnesium (Mg) and N deficiencies.
Magnesium (Mg)	Interveinal chlorosis on older leaves which proceeds to the younger leaves as the deficiency becomes more severe. The chlorotic interveinal yellow patches usually occur toward the center of the leaf with the margins being the last to turn yellow. Curling of leaves upward along margins.	High Mg can cause Ca deficiency.
Calcium (Ca)	Light green color on uneven chlorosis of young leaves. Brown or black scorching of new leaf tips and die-back of growing points. Growing points of stems and roots cease to develop. Poor root growth and roots short and thickened.	High Ca can cause Mg or Boron (B) deficiencies.
Sulfur (S)	Uniform chlorosis first appearing on new leaves.
Iron (Fe)	Interveinal chlorosis of new leaves followed by complete chlorosis and or bleaching of new leaves. Stunted growth.
Zinc (Zn)	Interveinal chlorosis of new leaves with some green next to veins. Short internodes and small leaves. Rosetting or whirling of leaves.
Manganese (Mn)	Interveinal chlorosis of new leaves with some green next to veins and later with grey or tan necrotic spots in chlorotic areas.
Copper (Cu)	Interveinal chlorosis of new leaves with tips and edges green, followed by veinal chlorosis. Leaves at the top of the plant wilt easily followed by chlorotic and necrotic areas in the leaves. Dieback of terminal shoots in trees.
Boron (B)	Death of terminal buds, causing lateral buds to develop and producing a ‘witches broom’ effect.	Symptoms develop as a yellow-tinted band around the leaf margins. The chlorotic zone becomes necrotic and gray, while the major portion of the leaf remains green.
Molybdenum (Mo)	Older leaves show interveinal chlorotic blotches, become cupped and thickened. Chlorosis continues upward to younger leaves as deficiency progresses.

Diagnosing Nutrient Deficiencies

EC and pH

pH and EC	symptoms	corrections
High salts EC	Small, thick, and dark colored leaves. Lower leaves may turn yellow and brown, especially on the leaf margins. Plants become stunted. Roots are less vigorous, may have brown leaves, and are likely to become diseased.	The solution to high salt levels is usually to “leach out” (apply excess water to wash through the container) the growing medium or potting mix (termed the “root substrate” throughout this book).
Low EC ( see specific nutrient deficiency)	“chlorosis” (yellowing, resulting from lack of the green chlorophyll pigment that is essential for photosynthesis). These symptoms can be seen on this geraniu Nitrogen, phosphorus, and other nutrients are being moved out of its lower leaves to provide N and P to the growing points. The plant is essentially cannibalizing itself, because it cannot get nutrients from the roots. Growth and flowering can be greatly reduced	The solution to low fertilizer levels is simply to apply more fertilizer. However, it is important to diagnose that nutrient levels are indeed low using an onsite soil test method and checking EC is below the adequate range
Substrate pH is too high (alkaline, pH values above 7 are “basic”)	e.g. Iron Symptoms to look for with iron deficiency at high pH: Chlorosis either in the entire leaf or between the darker leaf veins (“interveinal chlorosis”) Symptoms show up first in young leaves, in contrast to an overall low nutrient level, which may be in all or older leaves. With sensitive plants such as the calibrachoa above, growth is severely stunted and growing points turn white and even necrotic (the tissue dies).	Reduce excess limestone, or alkaline water, nitrate fertilizer’s basic effects. To correct a high pH problem, a combination of acid fertilizer (containing ammonium or urea nitrogen) and adding acid into the irrigation water to remove alkalinity can usually drop substrate-pH. Drenches (irrigations applied to the root substrate) with additional iron in a highly soluble form are also very effective at corr
Substrate pH is too low (acidity)	Mn and Fe toxicity (over availability) Chlorotic (yellow) or necrotic (brown and dead) spots or leaf margins in older leaves, In marigold, the symptoms appear as brown sandy-colored spots in older leaves	The most common causes of a drop in substratepH are insufficient limestone, a fertilizer high in ammonium or urea nitrogen, which has an acidic reaction. Some plant species such as geranium also tend to drop substrate-pH. An application of a liming material such as flowable limestone or potassium bicarbonate, and a change to a basic nitrate fertilizer are needed to raise substrate-pH

Fertilizer calculation one

	N	P	K
% weight	15	5	15


desired ppm = mg/L or mg/kg	hoal
( 1 liter oof water weighs 1 kg)	200	mg/L

		0.15 mg of N per mg og Fertilizer
	200 / 0.15 =
	1333.333333
		mg N / L / mg N/ mg F =
		mg F / L ) for desired 200 ppm)

		divide by 1000 to get grans
	1.333333333	g per L

		for 2000 L solution	2000

	2666.666667	grams F in 2000 L	1000
	2.666666667	kh of F in 2000 L

Fertilizer calculation two

1 gallon = 3.7854 L

	5000	gallon holding tank
		* 3.79
		3.79

	18,950.00	Liters storage tank

Calcium Nitrate		0.155
15.5 % N		mh N per Mg Ca Bitrate
	645.1612903	=100/0.155
desired ppm 100 )mg/L)	0.64516129	grams

	12,225.81	times storage tank
	12.23	kg

Two photosynthetic reaction centers are arranged in tandem in photosynthesis of algae and plants

Water is split by photosystem II

Photosystem II complex is very similar to the reaction center in purple bacteria

Mechanized agriculture usually necessitates the use of herbicides

The cytochrome-b6/f complex mediates electron transport between photosystem II and photosystem I Iron atoms in cytochromes and in iron-sulfur centers have a central function as redox carriers

The electron transport by the cytochrome-b6/f complex is coupled to a proton transport

The number of protons pumped through the cyt-b6/f complex can be doubled by a Q-cycle

Photosystem I reduces NADP

The light energy driving the cyclic electron transport of PS I is only utilized for the synthesis of ATP

In the absence of other acceptors electrons can be transferred from photosystem I to oxygen

Regulatory processes control the distribution of the captured photons between the two photosystems

Excess light energy is eliminated as heat

Photosynthesis is an electron transport process

The photosynthetic machinery is constructed from modules

Section 6.1 – An Introduction to Oxidation Reduction Reactions (10:27)

Section 6.2 – Oxidation Numbers (22:09)

Section 6.4 – Voltaic Cells (24:17)

Vides

o-Oxidation and reduction review from biological point-of-view | Biomolecules | MCAT | Khan Academy

A reductant and an oxidant are formed during photosynthesis

The basic structure of a photosynthetic reaction center has been resolved by X-ray structure analysis

X-ray structure analysis of the photosynthetic reaction center

The reaction center of Rhodopseudomonas viridis has a symmetric structure

playlist–

How does a reaction center function?

The use of energy from sunlight by photosynthesis is the basis of life on earth

Chlorophyll is the main photosynthetic pigment

Pigments capture energy from sunlight

The energy content of light depends on its wavelength

Light absorption excites the chlorophyll molecule

Video- Light-Harvesting: Photosynthetic Pigments and Exciton Transfer (Playlist photosynthesis)

An antenna is required to capture light

How is the excitation energy of the photons captured in the antennae and transferred to the reaction centers?

The function of an antenna is illustrated by the antenna of photosystem II

phycobilisome : Energy and Electron Transfer in Photosynthetic Organisms,

Photosynthesis: What are Photosystems? Resonance, Reaction Centers and Antenna Complex?

Figure 2.16 Absorption spectra of the phycobiliproteins phycoerythrin, phycocyanin, and allophycocyanin. For the sake of comparison

chlorophyll-a is shown

video-Visualizing Soil Properties: Water Infiltration

OSU Extension – Containers and Media for the Nursery

Growing Media for Container Production in a Greenhouse or Nursery

Media Chemical Properties

Cation Exchange Capacity (CEC)

Soluble Salts

Adjusting Media pH

Raising Media pH

Lowering Media pH

Managing Substrate EC

Disinfecting Media

Steam Pasteurization

Chemical Fumigation

Solarization

Media Physical Properties

Weight (Bulk Density)

lumetric Moisture Content (sometimes referred to as Water-Holding Capacity)

Graph-Electrical conductivity for various soil substrates

Porosity: Measuring Pore Space

Porosity (%) = 100 x Volume of water added to fill Pot with substrate / Volume of Pot

Air space = 100 x Volume of water drained / Volume of Pot

Water space (%) = 100 – Air space(%)

Table 1. The following table provides information about the fractional volume of pore space, air space, and water space in two soilless substrates:

Note AFP has different definitions- some have air + water + solid -100

Soil Electrical Conductivity

Factors Affecting

Relationship to Soil Function

Problems with Poor Soil EC Levels

Improving Soil EC

Measuring Soil EC

Monitoring electrical conductivity in soils and growing media

More References

Cation Exchange:

The reaction is as follows:

Cation Exchange Capacity (C.E.C.):

Factors affecting the Cation Exchange Capacity:

(i) Soil texture:

(ii) Organic matter content:

(iii) Amount and kind of clay:

(iv) pH:

Milliequivalents:

Percentage Base Saturation of Soils:

The cytochrome-b₆/f complex mediates electron transport between photosystem II and photosystem I Iron atoms in cytochromes and in iron-sulfur centers have a central function as redox carriers

The electron transport by the cytochrome-b₆/f complex is coupled to a proton transport

The number of protons pumped through the cyt-b₆/f complex can be doubled by a Q-cycle

Photosystem I reduces NADP^