What does the acronym 'PIN' stand for referring to PIN proteins in plants?

What does the acronym 'PIN' stand for referring to PIN proteins in plants?

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There are so called PIN proteins, or PIN-formed proteins, in plants. What does this acronym mean?

Wikipedia briefly explains the function of the protein but not the origin of the name. It's not explained even in the link-linked review paper, if I'm not missing something.

Like many genes and gene products, PIN proteins were named for a mutant phenotype and PIN is not actually an acronym; the source in your link does actually explain this (emphasis mine):

The significance and function of AtPIN1 was discovered through the phenotype generated by the loss-of-function mutation in the gene: mutant plants fail to develop floral organs properly and generate naked, pin-like inflorescences, which gave the name PIN-FORMED (PIN) to the family

Křeček, P., Skůpa, P., Libus, J., Naramoto, S., Tejos, R., Friml, J., & Zažímalová, E. (2009). The PIN-FORMED (PIN) protein family of auxin transporters. Genome biology, 10(12), 249.

It is not an acronym; it is an abbreviation of pinnulate (or pinnulated), which means having the pinnulae of a pinnate leaf properly developed.

Gene symbols written in all caps indicate the wild-type condition of the organism: If a gene is referred to as PIN1, it means its product (an auxin transporter) permits normal branching (or at lest does not contribute to malformations); when it is written in lower case, pin1, it indicates the gene has a variant associated with a mutant (non-pinnate) phenotype.

PIN1: nothing is broken

pin1: a single stalk without pinnulae (in the case of Arabidopsis)

If the mutant looks like a pin, it is a co-incidence (probably a mnemonic suggested by a English-speaking teacher). The original meaning derives from pinna, which refers to the normal appearance of leaves and flowers.

Oxidation Definition and Example in Chemistry

Two key types of chemical reactions are oxidation and reduction. Oxidation doesn't necessarily have anything to do with oxygen. Here's what it means and how it relates to reduction.

Key Takeaways: Oxidation in Chemistry

  • Oxidation occurs when an atom, molecule, or ion loses one or more electrons in a chemical reaction.
  • When oxidation occurs, the oxidation state of the chemical species increases.
  • Oxidation doesn't necessarily involve oxygen! Originally, the term was used when oxygen caused electron loss in a reaction. The modern definition is more general.


Post-segregational cell killing (PSK) is a widespread mechanism that aids several plasmids to maintain themselves in their bacterial hosts [1–4]. Operons containing genes for interacting toxin-antitoxin (T-A) pairs that are borne on these plasmids, are the basis for PSK. Typically, the first gene in these operons encodes a labile antitoxin, which also acts as a transcriptional regulator of the operon, while the second gene encodes a stable toxin. Usually, the antitoxin forms a physical complex with the toxin and neutralizes its action. A variation on this theme is seen in the form of the unstable anti-sense RNAs, which act as inhibitors of translation of the toxin mRNAs. If the plasmid is lost, the antitoxin is rapidly degraded while the stable toxin lingers on, killing cells that lack the plasmid. Thus, plasmids with systems for PSK cause their host cells to become addicted to them [1–4]. Additionally, several of these T-A systems are also found on prokaryotic chromosomes, where they may have alternative regulatory functions [5].

A systematic survey of such T-A operons and their mechanisms was presented in the seminal work of Gerdes in 2000 [6]. Subsequently, there have also been some important studies that have elucidated the biochemical details regarding the action of several toxins. One of these toxins, ParE, was shown to act as an inhibitor of the DNA gyrase, and it induced formation of DNA-gyrase covalent complexes, which could inhibit replication and damage the integrity of the chromosome [7]. In contrast, the RelE and Doc toxins were shown to be inhibitors of translation [5, 8]. More recently, it was demonstrated that the RelE protein cleaved transcripts associated with the ribosome, by specifically targeting codons associated with the ribosomal A-site [9]. RelE displays codon-specificity by showing highest preference for UAG among the stop codons and UCG and CAG among the sense codons [9]. Interestingly, this inhibition of translation by RelE is reversed by the transfer-messenger RNA (tmRNA), which acts as a regulator of protein stability in bacteria [10]. These studies have also suggested that the chromosomal versions of these antitoxin-toxin pairs could function as regulatory switches that control gene expression under poor growth conditions.

Although Gerdes proposed that all T-A operons could have a common origin [6], an objective evaluation of the evolutionary relationships of these proteins and the origin of these systems has not been conducted. The availability of a large number of prokaryotic genome sequences allows us to use a variety of computational approaches to address the problem of the origin and evolution of these systems. One approach, involving sensitive sequence searches using profile methods, allows the detection of distant relationships, which were hitherto not detected [11–13]. Additionally, it also enables objective evaluation of relationships, based on statistical significance of the detected similarities and multiple alignment-derived secondary structure predictions. A second approach involves the use of comparative genomics to detect conserved gene neighborhoods, gene or domain fusions, and to extract functional and evolutionary information from these contextual connections [14–18]. This approach is particular useful in the case of the prokaryotic PSK systems because of the strong coupling of the toxin and antitoxin genes in a single operon. Our objective in applying these analyses was to discover new functional connections that may not have been previously uncovered in experimental studies on these systems. Given the recent experimental results suggesting a specific role for these systems in the regulation of cellular responses to stress [9, 10, 19], we were also interested in identifying novel genomic versions of PSK-related systems with a wide phyletic distribution.

As a result of our analyses we were able to uncover several new T-A systems and establish an evolutionary relationship between them and the eukaryotic nonsense-mediated RNA degradation system. We also present evidence that the RelE and ParE families of toxins, despite their very distinct modes of action, have been ultimately derived from a common ancestor. Furthermore, we show that the Doc toxin defines a large family of enzymes that could potentially act on RNA and function as regulators of translation in both prokaryotes and eukaryotes.


Developmental biologists often conceptualize patterning mechanisms in uniform fields of cells, but in reality, positional information may be generated and translated in dynamic circumstances in which cells divide, grow, and change shape. These circumstances allow for unexpected feedbacks, making analysis a challenge. Rhizotaxis, the arrangement of lateral organs along plant roots, is a good example of a patterning process that occurs in a dynamic context. The mechanisms that regulate rhizotaxis have long remained a mystery, although the origin of lateral roots was described as early as 1888 [1]. In Arabidopsis, lateral roots arise from two files of pericycle cells that lie adjacent to the protoxylem [2], and the pattern of emerged lateral roots can be described in terms of the longitudinal spacing along a file and the left/right component (Figure 1A). The longitudinal pattern is variable and cannot be explained by mechanisms that require a fixed amount of time, distance, or number of pericycle cells between initiations [3]. There is, however, a strong tendency for lateral roots to arise on the outside, i.e., convex side, of the curve [4]. This tendency correlates with above-average auxin response at the proximal end of the meristematic zone (MZ) well before the first asymmetric divisions [5]. Lateral root formation can be induced by global increases in auxin content [6], and more specifically, by local activation of auxin synthesis in pericycle cells [7]. Mutations that render plants less sensitive to auxin reduce lateral root numbers [8,9]. Additionally, chemical or genetic inhibition of auxin transport can decrease lateral root density [10–12]. These observations indicate that lateral root formation is influenced by auxin, but they do not reveal the underlying mechanism. Here, we combine experimental and multilevel modeling approaches to unravel the molecular and biophysical mechanism that regulates rhizotactic patterning.

(A) Lateral roots are formed on the outside of curves in an alternating left/right rhythm (o indicates the outside, and i the inside of curve L indicates left, and R right, relative to the main axis of the root).

(B–D) Examples of root curvature resulting from gravitropic stimulation of different time intervals (B) 3 h (C) 4.5 h (D) never returned. Black asterisk (*) indicates the position of the emerged lateral root red arrowhead indicates the center of curve. Scale bar represents 500 μm.

(E) Lateral root formation correlates with the degree of root curvature resulting from gravitropic stimulation over various amounts of time. Symbols indicate time in the inverted position. Distance from the center of the curve to the nearest emerged lateral root is reported.

(F) Lateral root initiation is induced in manually curved roots. Left: location along the curve where lateral roots form is reported alongside the comparable position(s) for straight roots. Center of curve is defined as zero, and negative values are closer to the root tip, distal to the center of the curve. The curve was made 0.5 cm from the root tip. Right: percentage of lateral roots forming on each side of the main root. Sides are defined as inside and outside for curved roots, and left and right for straight roots, as shown in Figure 1A.

(G) DR5::GFP accumulates asymmetrically in the stele of manually curved roots. Solid red symbols indicate the outer half of the stele open black symbols indicate the inner half.

(H) Root curvature due to gravitropic response results in inverse asymmetric auxin distributions in the primary root MZ. DR5::vYFP (nuclear), PIN7:GFP (ER). Open arrowhead indicates the gravity vector during initial growth the solid arrowhead indicates the gravity vector during the period of inversion and the circled cross indicates the gravity vector directed into the plane during imaging. Scale bar represents 100 μm.

(I–L) Auxin response is enhanced locally in the pericycle and adjacent endodermal cell at a curve prior to the asymmetric cell division. Fluorescent markers as in (H). (I) 300 min, (J) 110 min, and (K) 10 min before, and (L) 10 min after the pericycle cell division. Arrows mark the location of the dividing nucleus. Scale bar represents 100 μm.


PIN1 Responds Slowly to Stem Decapitation

In our previously reported basic model for shoot branching control the auxin transport network across the stem is considered as a unitary system, with the plasma membrane accumulation of a generic PIN protein established and maintained by a canalization-based mechanism [33]. If this assumption is correct, PIN accumulation and polarization in stems should respond strongly to auxin flux. To test this hypothesis, we used isolated 2cm stem segments from basal primary inflorescence internodes of 5- or 6-wk-old plants expressing a well-characterized PIN1-GFP fusion protein from its native promoter, which complements the pin1 mutant [37]. This construct is expressed in the xylem parenchyma and cambial cells of the stem vascular bundles (stem anatomy is summarized in Fig 1), which are tissues classically associated with the PATS [20,38]. This expression pattern is identical to that of a different PIN1pro:PIN1-GFP line described previously [10,39].

A) Light micrograph of transverse section through the basal internode of a 6-wk-old Arabidopsis inflorescence stem. Vascular bundles are clearly visible as dark green triangles. Longitudinal sections used in this study were made across the center of the stem, between two vascular bundles, as indicated by the white line. B, C) Close up of cellular anatomy in a transverse section of an Arabidopsis vascular bundle and surrounding tissue. (C) is shaded to indicate tissue types. D) Light micrograph of longitudinal section through the basal internode of a 6-wk-old Arabidopsis inflorescence stem (along the type of transect indicated in A). Vascular bundles are visible as continuous white lines (indicated by white arrow heads). Inset shows the whole 2 cm segment. White box indicates the region shown in E, F. E,F) Close up of cellular anatomy in longitudinal section of an Arabidopsis vascular bundle and surrounding tissue. (F) is shaded to indicate tissue types. Unshaded/P = pith, purple/XP = xylem parenchyma, yellow/X = xylem, green/C = cambium, red/Ph = phloem, orange/E = epidermis, blue/IFF = interfascicular fibers.

The 2 cm segments were held vertically between two sections of agar (after [40]) through which different treatments could be applied. Firstly, we assessed how PIN1-GFP responds to the absence of any such treatments (“untreated”), approximating the effect of decapitation of intact plants. We reasoned that, since the rate of auxin transport in stems is typically measured as

6–10 mm/hour [41], any endogenous auxin present in these segments at the time of excision would be depleted within 4 hours. On the assumption that auxin flux maintains PIN1 polar localization, we therefore expected to observe rapid changes in PIN1 localization over this time-frame. However, we observed that PIN1 behavior is remarkably stable in this scenario. No obvious change in PIN1-GFP localization or expression occurred 4 hours after the segments were isolated (Fig 2A, 2E and 2I). These results suggest that either auxin depletion is not sufficient to trigger PIN1 endocytosis, or that auxin depletion has not occurred in these segments on the expected timescale.

PIN1-GFP expression in xylem parenchyma cells (see Fig 1) in longitudinally hand sectioned

2 cm basal inflorescence stem segments of 6-wk-old PIN1pro:PIN1-GFP plants. “Apical” and “basal” (at the left) refer to which end of the segment is being imaged. Numbers in the right hand corner indicate the number of segments/the number examined in which basal, polar PIN1 localization was seen in this treatment. Green signal indicates PIN1-GFP, red signal is chloroplast autofluorescence. A) Freshly harvested stem segments. B,D) Stem segments held vertically in Petri dishes between 2 agar blocks with 1 μM NAA supplied in the apical block for 1, 3, or 6 d, respectively. E–L) Stem segments held vertically in Petri dishes between 2 agar blocks with no hormone treatment (‘—‘) for 4 hours (E, I), 1 d (F, J), 3 d (G, K) or 6 d (H, L) at the apical end (E–H) or basal end (I–L) of the segment.

After 1 d, PIN1-GFP on the basal plasma membrane was markedly reduced or absent at the apical end of many segments (7/17, Fig 2F). However, at the same time point, there was still strong basally localized PIN1-GFP in the medial (not shown) and basal parts of the stem segments (Fig 2J), and this signal persisted for up to 6 d after isolation, although gradually weakening (Fig 2K and 2L). In contrast, when we added 1μM naphthalene acetic acid (NAA), an auxin analog, to the apical block of agar (simulating an intact shoot apex), we observed that PIN1-GFP remained strongly expressed and polarized to the basal membrane throughout the stem segment for up to 6 d (Fig 2A–2D). Thus, auxin clearly promotes ongoing PIN1 localization at the basal plasma membrane of xylem parenchyma cells, consistent with the hypothesis that the presence of PIN1 at this location 4 hours after decapitation results from slower-than-expected auxin depletion. This hypothesis is further supported by the strong divergence in PIN1 behavior along the 2 cm segments, with an apical to basal progression in PIN1 depletion over 6 d, suggesting that rather than simple rapid basipetal auxin depletion, auxin transport dynamics in the stem are more complex.

Stem Auxin Transport Kinetics Are Nonlinear

To test the idea that auxin levels in stem segments decline more slowly than we initially anticipated, we directly measured auxin levels in both the tissue and eluate of stem segments by gas chromatography-mass spectrometry (GC-MS). We found that freshly harvested basal stem segments contain on average 41 pg IAA per mg tissue (fresh weight) (standard deviation = 18, n = 4), equating to approximately 1,400 pg in a 20 mm segment (average mass

35mg), providing a benchmark for total IAA at t = 0 (Fig 3A). We then assayed, also by GC-MS, the auxin eluted from the basal end of freshly harvested stem segments over a time-course, by serially transferring the segments to fresh collection buffer. In this way, we collected auxin in the time intervals 0–0.5 hours, 0.5–1 hours, 1–2 hours, 2–4 hours, 4–8 hours, and 8–24 hours. Almost half the auxin was collected in the first two hours, consistent with the rapid auxin depletion we expected from previously documented transport rates for auxin (Fig 3A). However, elution of auxin continued over the next 22 hours of the experiment, leading to an average cumulative eluate at 24 hours of 1218pg (standard deviation 42, n = 4) suggesting that by t = 24 hours approximately 10% of the original auxin content remained undrained (Fig 3A). However, when we analyzed tissue auxin content of stem segments that had been allowed to drain for 24 hours in the same manner, we found an average of 17.6 pg/mg IAA (standard deviation = 4, n = 4) equivalent to approximately 600 pg of IAA per segment (43% of the t = 0 content), consistent with net synthesis of

400 pg IAA during the experiment. Taken together these data suggest that, in addition to a rapidly moving pool of auxin in the PATS, there may be a more slowly moving pool of auxin within stems and perhaps continued auxin synthesis. The combination of these factors appears to be sufficient to maintain PIN1 expression at the plasma membrane in the basal part of stem segments for many days following decapitation.

A) Endogenous auxin eluted from 2 cm stem segments from the basal inflorescence internode of 6-wk-old plants was quantified by GC-MS at different points post excision. The cumulative auxin collected (pg) is shown relative to time. The red line indicates the approximate free auxin content of the stem segments at t = 0. B) Distribution of radio-labelled IAA (measured as CPM) in 2 mm intervals of 24 mm long stem segments after a 10 min pulse of 5 μM radio-labelled IAA was supplied to the apical end of the segment. Stems were dissected and analyzed by scintillation 30 min (blue line), 60 min (red line) or 90 min (green line) after the application of the pulse n = 8 per time point, bars indicate standard error of the mean (s.e.m.). C) Distribution of radio-labelled IAA (measured as CPM) in 2 mm intervals of 24 mm long stem segments after a 10 min pulse of 5 μM radio-labelled IAA was supplied to the apical end of the segment. Stems were dissected and analyzed by scintillation 120 min (purple line), 150 min (light blue line) or 180 min (orange line) after the application of the pulse n = 8 per time point, bars indicate s.e.m. D) Auxin transport assay to measure cross-stem auxin movement. Two schemes were trialed (b–c and d–e), in both of which the apical end of 18 mm stem segments were dissected so that radio-labelled auxin could be supplied to only half of the stem (see illustration). Longitudinal incisions were made across the diameter of the stem (top image, red dotted line), to a depth of 5 mm, followed by a second transverse incision to remove half the stem. Control segments (a) were left intact. In the d–e scheme, the basal end of the segment was similarly treated at the start of the experiment to leave either half the stem directly beneath the site of auxin application (d) or diametrically opposite the site of application (e), while in the b–c scheme the basal end was left intact during the assay. The apical end of the stem segments was then immersed in 2μM 14C IAA for 6 hours. At the end of the assay, the basal 5 mm of stem was dissected from the stems. In the b–c scheme, the basal 5 mm was longitudinally bisected, to separate the tissue directly under the site of auxin application (b) from the tissue diametrically opposite (c). The amount of radio-label transported into the basal 5 mm in each of a, b, c, d, and e was then measured by scintillation. The graph shows the auxin transported in each of these dissections (measured as CPM), n = 16, bars indicate s.e.m.

These data suggest that auxin transport in the stem may have complex kinetics. To test this hypothesis, we developed a pulse assay in which stem segments were inverted with their apical ends immersed in a solution of radio-labelled auxin for 10 min before being transferred to tubes containing media without auxin. We then tracked the distribution of radio-label within the segments over time, by cutting the segments into 2 mm sections and quantifying the radio-label present in each section. After 30 min, there was a relatively tight peak of radio-label towards the apical end of the segment, at a position consistent with most auxin molecules being transported at the commonly quoted rate of approximately 1 cm/h however, a proportion of the auxin molecules in this assay moved substantially faster (Fig 3B). At subsequent time points (60 and 90 min), the auxin distribution became broader and shallower, and a distinct peak was difficult to discern (Fig 3B). Treatment with the auxin transport inhibitor 1-N-naphthylphthalamic acid (NPA) blocked the movement of auxin through these segments, showing that these profiles arose from active auxin transport (S1 Fig). From 120–180 min the radio-label gradually accumulated at the basal end of the stem, leaving only a very small residue across the rest of the stem (Fig 3C).

This rather diffuse pattern of auxin distribution is not consistent with auxin moving solely through the high conductance PATS. We hypothesized that this pattern arises because there is significant exchange between the PATS and the surrounding tissues [42]. The population of radio-labelled auxin molecules in this assay therefore does not move in a simple linear fashion though the transport stream, but is gradually spread along the stem and eventually collects at the base, moving on average more slowly than anticipated, but with a very large variance in the movement rate of the labeled auxin molecules.

Auxin Moves across Stems

We have previously observed that two consecutive buds on an isolated stem segment can inhibit each other’s growth, despite connecting, and therefore exporting auxin, into different vascular bundles in the main stem [36]. In the context of slower and more complex than expected movement of auxin along the stem, we hypothesized that this inhibition might arise from auxin movement across the stem. To assess whether cross-stem auxin movement occurs we developed a cross-stem transport assay (Fig 3D), based on a modification of our basic bulk auxin transport assay in which stem segments are treated apically with radio-labelled auxin solution for 6 hours. For the cross-stem assay, radio-labelled auxin was supplied apically to only half the stem, and was then measured basally in either the same half, or the opposite half of the stem. If auxin moves strictly basipetally through the transport stream in each vascular bundle, there should be little radio-label collected in the opposite half, since within a single internode there is no vascular connection between the site of auxin application and collection [43]. However, in line with our hypothesis, a significant amount of auxin was detected on the opposite side of the stem to the site of auxin supply (Fig 3D). Taken together, these data suggest that, in addition to the classical PATS that is associated with vascular bundles, there is also appreciable auxin transport through a wide variety of tissues in the stem.

PIN1 Expression in the Stem Is Auxin Inducible but Highly Cell Type-Specific

The observed slow depletion of auxin suggests that the slow depletion of PIN1 (Fig 2) could reflect a canalization-like mechanism, with auxin flux being required to maintain PIN1 polarity. A further prediction of the canalization hypothesis is that auxin can induce expression of auxin transporters in naïve tissue, and indeed expression of the PIN1 gene has previously been shown to be auxin-inducible in root and shoot apical meristems [44,45]. We therefore investigated whether the observed changes in PIN1 expression in stem segments could be explained by changes in PIN1 transcription. Using quantitative PCR, we analyzed transcription of PIN1 in stem segments treated with apically applied 1μM NAA for 3 d, or left untreated for 3 d, compared to equivalent fresh stem segments. We found that PIN1 transcription is induced approximately 6-fold by auxin treatment, and reduced approximately 10-fold in the absence of auxin (Fig 4A) expression of a control auxin-inducible gene, MORE AXILLARY GROWTH4 (MAX4), behaved as expected in response to decapitation/auxin treatment [28]. However, when we analyzed GFP accumulation patterns in PIN1pro:PIN1-GFP plants in auxin treated segments, we did not observe any obvious change in the PIN1 expression pattern, even in very young stem segments taken immediately post-bolting. Cell files either expressed PIN1 or they did not, regardless of auxin treatment (Fig 4C–4E). This suggests that PIN1 expression in the stem is auxin inducible but highly cell-type specific, and/or there is cell-type specific post-transcriptional regulation.

Spatio-temporal regulation of auxin signalling capacities also contributes to the SAM patterning

The action of auxin in a patterning process depends not only on the distribution of auxin in a tissue but also on the cellular capacities to sense auxin and trigger specific transcriptional responses. Protein–protein interactions between the Aux/IAA and auxin response factors (ARF) transcriptional regulators are central to auxin signalling. There are 29 Aux/IAA genes that encode mostly short-lived repressors of auxin-induced transcription. The 23 ARF proteins can be either activators (the five Q-rich ARFs) or repressors of transcription. Aux/IAA and ARF proteins are able to form homo- and heterodimers both within and between the families. The instability of the Aux/IAAs is intrinsic to the so-called domain II, which interacts directly with SCF-like ubiquitin protein ligases harbouring TIR1 or one of the three related AFB F-box proteins ( Dharmasiri et al., 2005a, b Kepinski and Leyser, 2005 Tan et al., 2007). TIR1 and the AFBs act as auxin co-receptors together with the Aux/IAAs ( Calderón-Villalobos et al., 2012), and their activation leads to auxin-dependent degradation of Aux/IAAs. A model for auxin transduction is that Aux/IAAs dimerize with the ARF activators. These complexes bind to auxin-inducible genes, thus preventing transcription. By promoting the degradation of the Aux/IAAs, auxin would allow the ARFs to activate transcription. Most Aux/IAAs are themselves targets of the ARFs, thus establishing a negative-feedback loop.

Treatment of pin1 meristems with exogenous auxin demonstrated that all the cells at the periphery of the meristem are competent to initiate organs in response to auxin ( Reinhardt et al., 2000). However, auxin cannot trigger organogenesis at the centre of the meristem. Mutation in MONOPTEROS/ARF5 (MP/ARF5) induces a pin-like phenotype and also blocks the ability of the PZ cells to respond to exogenous auxin ( Hardtke and Berleth, 1998 Reinhardt et al., 2003). MP/ARF5 has been shown to exert its function in organogenesis, in part, by directly inducing, in an auxin–dependent manner, the expression of LFY, ANT, and ANT-like 6 (AIL6) TFs ( Yamaguchi et al., 2013). As MP/ARF5 is only expressed at the meristem periphery ( Hardtke and Berleth, 1998 Yamaguchi et al., 2013), the competence for organ initiation at the periphery of the meristem thus depends, at least in part, on a spatial modulation of auxin signal transduction. Using mostly in situ hybridization, a recent study identified TIR1, AFB1, and AFB5 and over 20 Aux/IAAs and ARFs as the effectors of auxin perception and signalling in the SAM. This analysis also demonstrated that most of these regulators are expressed differentially in the SAM, with a low expression at the centre of the SAM and high expression in the PZ. Notably, both ARF repressor and ARF activator expression followed this general trend. A full Aux/IAA–ARF interactome was obtained using a yeast two-hybrid approach, and knowledge on the topology of the Aux/IAA–ARF auxin signalling pathway was used to develop a mathematical model of the control of gene transcription by auxin in the SAM. This model predicted a role for the Aux/IAA–ARF pathway in creating a differential sensitivity to auxin between the centre and the periphery of the SAM, together with a capacity to buffer fluctuations in the auxin signal to stabilize the transcriptional response. These two predictions could be validated by analysing the differences between the spatio-temporal patterns of fluorescence of DII-VENUS and DR5 ( Vernoux et al., 2011).

This work shows that the spatial distribution of ARF genes in the SAM is essential to restrict high transcriptional responses to auxin (including Aux/IAA expression) at the periphery of the meristem and is essential to the definition of the two main functional domains of the SAM, the CZ and the PZ. In addition, the role of the signalling pathway in stabilizing transcriptional responses to auxin probably contributes to the robustness of patterning at the SAM and thus of phyllotaxis ( Vernoux et al., 2011). The dynamics of morphogenesis at the SAM thus appear to result from a dynamic integration of both spatial distribution of the auxin signal and local signalling capacities in the spatial control of morphogenesis ( Fig. 1), similarly to what has been observed recently for morphogen-driven developmental patterning in animal systems ( Kicheva et al., 2012).

Multiple feedback loops acting across different scales drive auxin-driven organ formation at the SAM. Auxin accumulation in cells at specific locations of the PZ in the SAM triggers both TIR1/AFB- and ABP1-mediated signalling. The activation of cellular auxin responses promotes: (1) PIN1-mediated auxin efflux that will in turn influence polar auxin transport patterns in the SAM and (2) the level of auxin biosynthesis in the SAM. These will in turn feed back on the patterns of auxin accumulation amplifying the local growth responses that promote organ formation.

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Wetlands Decoder Table Download

The database table in this download provides a crosswalk from U.S. Fish and Wildlife Service, National Wetlands Inventory (NWI) wetlands data, as defined by the Federal Wetland Mapping Standard, to the complete wetland definitions, as defined by the Federal Wetlands Classification Standard. The table can be joined with the NWI wetlands data using the 'Attribute' field. This will provide users with a full wetland or deepwater habitat description for each polygon. The NWI dataset and associated tables are updated on a biannual basis, typically in October and May. To ensure you have the most up to date information, please refer to the published date in the metadata, and download new data and tables regularly.

NWI Code Definitions Download Package - Last updated: May 2021

Phosphorus (P)

Phosphorus also plays a role in an array of functions necessary for healthy plant growth, contributing to structural strength, crop quality, seed production, and more. Phosphorus also encourages the growth of roots, promotes blooming, and is essential in DNA.

The transformation of solar energy into usable compounds is also largely possible because of phosphorus.

Sources of Phosphorus

Like nitrogen, phosphorus in NPK fertilizer can come from both organic and inorganic sources:

Common Inorganic Sources of P in NPK Blends

The primary source of inorganic phosphorus is phosphate rock. Crushed phosphate rock can be applied to soils directly, but it is much more effective if processed to be more readily available for plant uptake.

Common Organic Sources of P in NPK Blends


Our P. veris genome assembly exemplifies the power of high-throughput DNA sequencing technologies and establishes a benchmark for the rapid de novo assembly of a highly heterozygous, non-model plant with a moderately sized genome (that is, 479.22 Mb). We believe that the primary strength of our sequencing strategy lies in the diversity of sequence libraries and sequencing platforms we have employed. Our study suggests great promise in the application of PacBio long-read data for the improvement of de novo genome assemblies, and it is possible that, with the direct incorporation of the raw PacBio sequences in the assembly process, even more information could be extracted from long reads. Currently such integration is strongly limited by the a priori correction of raw PacBio reads, which significantly reduces the quantity of usable data resulting from a PacBio run (G. Russo, personal observation). In the case of large eukaryotic genomes, this translates into the need for a largely unfeasible sequencing throughput.

Using our de novo genome assembly coupled with RNAseq data from flower buds, we have identified 113 genes that show significant morph-specific differential expression in both P. veris and P. vulgaris. Functional analysis of the list of candidate genes has revealed clusters of GO terms related to extracellular processes, cell growth and organization, and development of reproductive structures. Furthermore, our genome assembly shows that the B-function MADS box gene GLOBOSA has been duplicated in Primula, and we find that one of these copies (PveGLO2) is silenced in L-morph flower buds, but we still do not know if PveGLO2 is linked to the S-locus, and thus a suitable candidate gene for morph-specific floral development. Future work toward characterizing this candidate gene could involve resequencing both L- and S-morph plants to evaluate the presence of this gene in both morphs and identify morph-specific SNPs that can be tested for S-locus linkage in a mapping population.

Employing a bulk segregant analysis followed by high-resolution mapping, we identify six loci on four genome scaffolds that are tightly linked to the S-locus in P. veris. When examining these S-linked genome scaffolds as well as genome scaffolds carrying genes previously identified as linked to the S-locus, we find elevated heterozygosity in S-morph versus L-morph coding sequences, consistent with theoretical predictions. Future work toward defining the recombinational limits and genetic composition of the S-locus would benefit greatly from the construction of a linkage map using genetic markers anchored within genome scaffolds. Beyond characterizing the Primula S-locus, our de novo genome sequence will prove to be a valuable resource for marker design and the analysis of population genomic and phylogenomic data. The genomic resources presented here thus represent a significant leap forward in the development of P. veris and P. vulgaris as models in the study of distyly, climatic adaptation, and speciation genetics.

Practice with the Codon Chart

Another great way to increase your knowledge of protein synthesis and better prepare for protein synthesis worksheets is to practice with the codon chart. You can find the solutions in parenthesis after the example:

  1. CUU-CGU-AAU-UGG-AAG (leu-arg-asn-trp-lys)
  2. ACU-ACA-AGU-UGC-UUU (thr-thr-ser-cys-phe)
  3. AAC-AAG-GUC-GUC-AGG (asn-lys-val-ile-arg)

Protein synthesis is a complex, highly tuned process that enables life to flourish. Understanding it, from the DNA to the RNA to the amino acids, gives us a better appreciation for life itself. Use our protein synthesis worksheet practice questions to help you learn the ins and outs of protein synthesis and remember the informaion.

Watch the video: DNA μέρος 2ο: Ας φτιάξουμε πρωτεΐνες! (July 2022).


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