Comparison between Monoclonal antibody vs. Aptamer:
\r\n\tThe major pathogenetic mechanisms resulting from RAAS overactivity include activation of the sympathetic nervous system, endothelial dysfunction, proinflammatory, and procoagulant states.
\r\n\tEmerging from basic science evidence, major clinical trials established the beneficial effects of inhibitors of the different components of RAAS such as angiotensin-converting enzyme (ACE) inhibitors, angiotensin receptor blockers (ARBs), aldosterone antagonists. These effects range from treatment of hypertension, diabetic nephropathy, CHF, as well as improvement of outcomes after myocardial infarction and improvement in glucose homeostasis and prevention of type 2 diabetes with some agents.
\r\n\tIn this book, written by a world-renowned scholar, we will address the major concepts and topics related to RAAS activation including the pathogenetic mechanisms underlying the deleterious effects of activated RAAS and the role of local tissue RAAS in various organ systems such as the heart and vasculature, the skeletal muscle, adipose tissues, pancreas and the angiotensinergic pathways in the brain. Cutting-edge information is provided that will address the need for a wide range of readers including a medical student, clinical practitioner, and basic science investigators alike. This book will be bridging the gap between basic science and clinical practice regarding the RAAS system, which is imminently critical and highly relevant to the practice of medicine.
\r\n\r\n\tFinally, with data emerging from the COVID-19 pandemic indicating overrepresentation of people with diseases associated with RAAS activation such as hypertension, chronic kidney disease, and diabetes, the role of RAAS activation and RAAS inhibition in the pathogenesis and clinical outcomes in COVID-19 has garnered a great deal of interest. In this book, we will dedicate a chapter addressing this topical and highly critical subject.
\r\n\t
Phloem is the vascular plant tissue responsible for the transport and distribution of sugars produced by the photosynthesis. Since the plant is a continuum, phloem will be found in the external part of root cylinders (Figure 1a), in the stem vascular bundles (Figure 1b) and in the abaxial part of the venations of every single leaf (Figure 1c). While the most common is to have the phloem external to the xylem in roots and stems and abaxial in leaves, some exceptions exist and are usually taxon specific. The phloem found in the inside is named internal or intraxylary phloem (Figure 1b).
Location of the primary phloem in different organs and its cell composition. (a) Ranunculus acris (Ranunculaceae). Root transverse section (TS), exarch structure, six strands of primary phloem alternating with the six protoxylem poles. (b) Bicollateral vascular bundle of a squash, Cucurbita pepo (Cucurbitaceae) TS. On the top is the external phloem, and on the bottom is the intraxylary or internal phloem. (Picture credit to Solange Mazzoni Viveiros). (c) Detail of the leaf midrib vascular cylinder of Tetrapterys mucronata (Malpighiaceae) showing primary xylem on the top and primary phloem on the bottom. (Picture credit to Leyde Nayane Nunes). (d) Detail of (b), showing the protophloem on top and the metaxylem on the bottom. ep, external phloem; ip, intraxylary phloem; mp, metaphloem; p, parenchyma cell; pp, primary phloem; ptp, protophloem; px, primary xylem. Scalebars: a, c, d = 50 μm, b = 130 μm.
As a constitutive tissue in the plant body, phloem functions extrapolate its main function of sugar transport, including transport of signalizing molecules such as mRNAs, hormones, defenses from biotic and abiotic agents, sustenance of the organs, gas exchange, and storage of many ergastic materials, such as starch, calcium oxalate crystals, and tannins. Parenchymatic cells of the phloem can also give rise to new meristems, such as the phellogen or cork cambium. All vascular plants have phloem, which typically includes specialized living conducting cells named sieve elements whose nucleus, ribosomes, and other organelles degenerate during maturation, making sugar transport more efficient. The life and function of these cells will then rely on closely associated parenchyma cells which support the physiological functions of these sieve elements [1]. Although typical phloem is exclusive of vascular plants, rudimentary phloem-like conducting cells are present also in other lineages, such as the bryophyte leptoids, and even outside the plant kingdom, as the trumpet cells of the kelps and phaeophycean algae [2]. The primary phloem derives from the embryo and the apical meristem procambium throughout the life of the plant or from the cambium, in plants with secondary growth.
The phloem is a complex tissue and is formed typically by three cell types, the sieve elements, the parenchyma cells, and the sclerenchyma cells (Figure 2a–d). Sclerenchyma cells might sometimes be absent in primary and/or secondary phloem. The presence, quantities, and arrangements of these cell types in the tissue commonly vary and may be taxonomic informative [3, 4]. Lists depicting these variations in all phloem cell types are of ultimate importance for complete bark descriptions [5]. What follows is a description of these three major cell types in the phloem.
General aspects of the secondary phloem. (a) Composition of the secondary phloem of Luehea divaricata (Malvaceae) TS, showing sieve tube elements (se) in clusters, axial parenchyma cells (p), fiber clusters (f), and rays (r). (b) Longitudinal tangential section (LT) of Cordia caffra (Boraginaceae) showing sieve tube element (se), companion cells (arrow), multiseriate ray (r), and axial parenchyma (p). Note callose staining with resorcin blue evidencing the slightly inclined simple sieve plates. Note also the P-protein (asterisk) next to the sieve plate. (c) LT of the secondary phloem of Castanea dentata (Fagaceae) showing sieve tube elements (se) with inclined, compound sieve plates and numerous lateral sieve areas of narrower pores, unicellular rays (r), and axial parenchyma (p). (d) TS of Talauma sp. (Magnoliaceae) showing sieve tube elements in clusters, with conspicuous nacreous walls, parenchyma cells (p), clusters of fibers (f), and rays (r). (e) Secondary phloem of maple, Acer saccharum (Sapindaceae), showing the conducting phloem (cp), where sieve tubes and companion cells are turgid, and the nonconducting phloem (ncp), with collapsed sieve tubes. (f) Secondary phloem of Carya cordiformis (Juglandaceae) showing a phloem formed by a background of fibers where solitary to multiple of two sieve tubes are scattered, with sieve-tube-centric and diffuse-in-aggregate axial parenchyma. Note that no collapse is seen in the nonconducting phloem of Carya. c, cambium; sx, secondary xylem. Scalebars: a = 100 μm, b-d = 50 μm, e, f = 200 μm.
Sieve element is a general term that encompasses all conducting cells of the phloem, both sieve cells and sieve tube elements [1, 6]. The name sieve derives from the strainer appearance given to the cells by the presence of numerous pores crossing their bodies (Figure 2c). These pores are specialized plasmodesmata of wider diameter, and the sieve areas are basically specialized primary pit fields [7]. The sieve pores are usually lined up with callose, which were shown to be related with the formation of the sieve pores in angiosperms, although not in gymnosperms [8]. Large amounts of callose deposit in the sieve areas also when the sieve element loses conductivity, suffers injury, or becomes dormant. Callose in gymnosperms is typically wound callose [8]. Callose can be easily detected with aniline blue under fluorescence or resorcin blue [9] (Figure 2b and c).
Sieve elements have only primary walls, but sometimes this wall can be very thick receiving the name of nacreous walls (Figure 2d) [10] and can be present in all major vascular plant lineages [1]. Nacreous walls can be very thick, and some authors have proposed they would be secondary walls [1, 8]. Nacreous walls can almost occlude the entire lumen of the sieve element (Figure 2d); hence, its presence needs to be considered in experiments of sugar translocation. Such thick walls might be related to resistance to high turgor pressures within the sieve elements. Nacreous walls seem to have a strong phylogenetic signal and are much more common in some families, such as Annonaceae, Calycanthaceae, and Magnoliaceae [10].
There are basically two types of sieve elements: sieve cells and sieve tube elements. The sieve tube elements are distinguished by the presence of sieve plates, that is, sieve areas with wider and more abundant sieve pores, usually in both extreme ends of the cells, while sieve cells lack sieve plates [1, 6, 8]. A group of connected sieve tube elements form a sieve tube [8]. According to this concept, lycophytes and ferns have sieve cells [1]. However, because of the many differences in the morphology and distribution of protoplasm organelles and chemical substances between the sieve elements of gymnosperms and vascular cryptogams, Evert [8] suggests the use of “sieve cell” as exclusive to the gymnosperms, leaving the more general term “sieve element” to the lycophytes and ferns.
The longevity of sieve elements varies. In many species it is functional for just one growth season, while for other species they can be functional a couple of years, or in the case of plants that lack secondary growth, they will be living for the entire plant life spam. Palm trees would perhaps be the plants with the oldest conducting sieve tube elements, since some reach 200 years [11]. In other plants, on the other hand, the sieve elements collapse a few cells away from the vascular cambium, corresponding to a fraction of the mm. In a mature tree, most of the secondary phloem will generally be composed of sieve elements no longer conducting. This region is called nonconducting phloem, in opposition to the area where sieve elements are turgid and conducting, called conducting phloem [5, 8] (Figure 2e and f). The term collapsed and noncollapsed phloem and functional and nonfunctional phloem are not recommended, since in some plants the nonconducting phloem keeps its sieve elements intact (Figure 2f), and although large parts of the phloem may not be conducting, the tissue as a whole is certainly still functioning in storage, protection, and even dividing or giving rise to new meristems, such as the phellogen and the dilatation meristem of some rays [5, 8].
Sieve cells are typically very elongated cells with tapering ends (Figure 3b), which lack sieve plates, that is, lack an area in the sieve element where the pores are of a wider diameter. Even though the sieve areas may be more abundant in the terminal parts of the sieve cells, the pores in these terminal areas are of the same diameter as those of the lateral areas of the sieve element. Sieve cells lack P-protein in all stages of development. The sustenance of the sieve cells is carried by specialized parenchyma cells in close contact with the sieve elements, with numerous plasmodesmata, which maintain the physiological functioning of the sieve cells, including the loading and unloading of photosynthates. These cells are known either as albuminous cells or Strasburger cells. The name albuminous was initially coined given the proteinaceous appearance of these cell’s contents. However, because the high protein content is not always present, the name Strasburger cell, paying tribute to its discoverer Erns Strasburger, is recommended over albuminous cells [5, 12]. Strasburger cells in the secondary phloem can be either axial parenchyma cells, as is common in Ephedra [13], or ray parenchyma cells, as is common in the conifers (Figure 3c) [14]. More commonly, the most conspicuous Strasburger cells in conifers are the marginal ray cells which are elongated (Figure 3c) and have a larger number of symplastic contact with the sieve cells [14]. Sometimes declining axial parenchyma cells also acts as Strasburger cells in Pinus [14]. The only reliable character to distinguish a Strasburger cell from an ordinary cell is the presence of conspicuous connections [14]. In the primary phloem, parenchyma cells next to the sieve cells are those which act as Strasburger cells.
The secondary phloem of conifers. (a) Transverse section of the secondary phloem of Sequoia sempervirens (Cupressaceae) showing alternating tangential bands of sieve cells, axial parenchyma, and fibers, interrupted by uniseriate rays. (b). Longitudinal radial section (LR) of the secondary phloem of Sequoia sempervirens (Cupressaceae) showing alternating tangential bands of sieve cells, axial parenchyma, and fibers, interrupted by uniseriate rays. Sieve pores distributed across the walls of long sieve cells. (c). LR section of Pinus strobus (Pinaceae) showing the elongated marginal ray cells in close contact with the sieve cells. These are the Strasburger cells. f and rectangular symbol = fibers, s and * = the sieve cells, p and dot = axial parenchyma cells rich in tannins. Scalebars: a, b = 100 μm, c = 50 μm.
A synapomorphy of the angiosperms is the presence of sieve tube elements and companion cells, both sister cells derived from the asymmetrical division of a single mother cell. In some instances, these mother cells can divide many times, creating assemblages of sieve tube elements and parenchyma cells ontogenetically related [15]. Sieve tube elements have specialized areas in the terminal parts of the sieve elements in which a sieve plate is present (Figures 2b and c). Within the sieve plate, the pores are much wider than those of the lateral sieve areas, evidencing a specialization of these areas for conduction [16]. In Cucurbita, the pores in the sieve plate have up to 10 μm in diameter, while the pores in the lateral sieve areas are of about 0.1 μm [7, 17]. The protoplast of sieve tube elements contain a specific constitutive protein called P-protein (P from phloem, also known as slime; Figure 2b), which in some taxa (e.g., Leguminosae) is nondispersive and can be seen as coagula inside of the sieve element [18].
Even in lineages of angiosperms where vessels were lost and tracheids re-evolved, such as Winteraceae in the Magnoliids and Trochodendraceae in the eudicots, sieve elements and companion cells are present [19], suggesting the independent evolution of these two plant vascular tissues derived from the same meristem initials. Since the sieve tube element loses its nucleus and ribosomes, the companion cell is the cell responsible for the metabolic life of the sieve elements, including the transport of carbohydrates in and out the sieve elements [7]. Companion cells may be arranged in vertical strands, with two to more cells (Figure 2b). Other parenchyma cells around the sieve tube integrate with the companion cells and can also act in this matter [7]. Typically, the cells closely related with the sieve tube elements die at the same time as the sieve element loses conductivity.
Sieve tube elements vary morphologically. The sieve plates can be transverse to slightly inclined (Figure 2b) or very inclined (Figure 2c) and contain a single sieve area (Figure 2b) or many (Figure 2c). When one sieve area is present, the sieve plate is named simple sieve plate, while when two to many are present, the sieve plates are called compound sieve plates. Compound sieve plates typically occur in sieve tube elements with inclined to very inclined sieve plates (Figure 2c). In addition, sieve elements with compound sieve plates are typically longer than those with simple sieve plates. Evolution to sieve elements of both sieve area types has been recorded in certain lineages, such as in Arecaceae, Bignoniaceae, and Leguminosae [5, 20], and to the present it is not still clear why the evolution of distinct morphologies would be or not beneficial. The only clear pattern is that compound sieve plates appear in long sieve elements [1], and phloem with a lot of fibers generally has compound sieve plates [20].
In the primary phloem, just one type of parenchyma is present and typically intermingles with the sieve elements (Figure 1d). In the secondary structure, there are two types of parenchyma: axial parenchyma and ray parenchyma (Figures 2b, c, 3b, c), derived, respectively, from the fusiform and ray initials of the cambium.
The axial parenchyma in conifers commonly is arranged in concentric, alternating layers (Figure 3a and b). These parenchyma cells contain a lot of phenolic substances, which were viewed as a defense mechanism against bark attackers [21]. In Gnetales, the phloem axial parenchyma appears to be intermingling with the sieve cells (Figure 4a) [22]. Some of these axial parenchyma cells act as Strasburger cells [13].
Phloem axial parenchyma distribution in secondary phloem. (a) Ephedra tweediana (Ephedraceae) TS, showing sieve cells interspersed by axial parenchyma cells (arrows). Six to five cells away from the cambium, the sieve cells already lose conductivity and collapse with axial parenchyma cells enlarging (top arrow). (b) Lannea discolor (Anacardiaceae) TS showing axial parenchyma with tannins arranged in narrow bands (arrows). There are also other parenchyma cells with less content dispersed in the phloem. Note also the fibers in concentric bands. (c) Robinia pseudoacacia (Leguminosae) TS showing bands of axial parenchyma associated with the fiber bands and sieve tube elements in clusters with simple sieve plates staining with resorcin blue. (d) Fridericia nigrescens (Bignoniaceae) TS with sieve tubes surrounded by sieve-tube-centric axial parenchyma. The tissue background corresponds to the fibers. c, cambium; sx, secondary xylem; c, cambium; sp, secondary phloem; sx, secondary xylem. Scalebars: a = 50 μm; b, d = 200 μm; c = 100 μm.
In angiosperms, the distribution of the axial phloem parenchyma is more varied, and it may appear as a background tissue where other cells are dispersed or may be in bands (Figure 4b and c) and radial rows or sieve-tube-centric (Figure 4d) [5, 20]. The distribution of axial phloem parenchyma is commonly related to the abundance of fibers or sclereids. In species with more fibers, it is common to have a more organized arrangement of the parenchyma. For example, in Robinia pseudoacacia (Leguminosae) there are parenchyma bands in either side of the concentric fiber bands (Figure 4c). When very large quantities of sclerenchyma are present, such as in the secondary phloem of Carya (Juglandaceae) or in Fridericia, Tanaecium, Tynanthus, and Xylophragma (Bignoniaceae), the sieve-tube-centric parenchyma appears (Figure 4c) and, as the name suggests, is surrounding the sieve tubes [8, 20, 23].
Although collectively described and referred to as axial phloem parenchyma, it is important to note that in many plants there will be distinct groups of phloem parenchyma within the phloem with quite different ergastic contents and therefore presumed different functions. Some of these specialized parenchyma cells may be considered secretory structures. Within a single plant, it is not uncommon that while some cells have crystals (especially when in contact with sclerenchyma), others have tannins, starch, and other substances. In apple trees (Malus domestica, Rosaceae) three types of axial parenchyma have been recorded: (1) crystal-bearing cells, (2) tannin- and starch-containing cells, and (3) those with no tannin or starch, which integrate with the companion cells [15].
Within bands of axial parenchyma, canals with a clear epithelium may be formed in many plant groups such as Pinaceae, Anacardiaceae, Apiales, a feature with strong phylogenetic signal. Some phloem parenchyma cells also act in the sustenance and support of the sieve elements, even when not derived from the same mother cell [7]. In longitudinal section, the axial phloem parenchyma may appear fusiform (not segmented) or in two up to several cells per strand [5].
While the phloem ages and moves away from the cambium, its structure dramatically change, and typically axial parenchyma cells enlarge (Figures 4a and b, 6c), divide, and store more ergastic contents toward the nonconducting phloem. In plants with low fiber content, the dilatation undergone by the parenchyma cells typically provokes the collapse of the sieve elements. The axial parenchyma in the nonconducting phloem can dedifferentiate and give rise to new lateral meristems. In plants with multiple periderms, typically new phellogens are formed within the secondary phloem, compacting within the multiple periderms large quantities of dead, suberized phloem. In plants with variant secondary growth, especially lianas, new cambia might differentiate from axial phloem parenchyma cells [24]. In the Asian Tetrastigma (Vitaceae), new cambia were recorded differentiating from primary phloem parenchyma cells [25].
Sclerenchymatic cells are those with thick secondary walls, commonly lignified. Sclerenchyma can be present or not in the phloem, and when present it typically gives structure to the tissue. For instance, a phloem with concentric layers of sclerenchyma cells is called stratified (Figures 2e, 3a, and 4c) [5]—not to be confused with storied, regarding the organization of the elements in tangential section. In Leguminosae, bands of phloem are associated to the concentric fiber bands (Figure 4c).
Older phloem shows more sclerification than younger phloem, and the sclerenchyma may also act as a barrier to bark attackers [21]. The sclerenchyma is typically divided in two categories: fibers and sclereids. These cell types differ mainly in form and size, but origin has also been used to distinguish them [26].
Fibers are long and slender cells, derived from meristems, the fiber primordia [1, 26, 27]. In the primary phloem, fiber caps are sometimes found in association with the protophloem (Figure 5a) and are named protophloem fibers. Since only an ontogenetic study can evidence whether these fibers indeed differentiate within the protophloem, a term coined in the nineteenth century German and American literature, pericyclic fibers, has been recommended to be used instead of primary phloem fibers or perivascular fibers [5]. In the monocotyledons, fibers are commonly an important component of the vascular bundles (Figure 5b–d). Commonly these fibers are associated with the phloem (Figure 5b), but they might also be associated with the xylem (Figure 5c) or be central in the vascular bundle (Figure 5d). These fibers are not, however, understood as part of either phloem or xylem; although they are of vascular nature, they differentiate directly from procambium.
Vascular fibers associated to eudicot and monocot primary structure. (a) Pericyclic fiber cap (fc) and primary phloem (pp) in Perianthomega vellozoi (Bignoniaceae). Secondary phloem (sp) beginning to be produced. Vascular bundles in monocotyledons. (b) Vascular bundle in the climber Calamus manan (Arecaceae) with fibers toward the phloem side. Phloem in two strands around a wide metaxylem vessel. (c) Vascular bundle of Vellozia alata (Velloziaceae), with fiber cap toward the xylem side. Phloem on the top side of the picture. (Picture credit to Marina Blanco Cattai). (d) Amphivasal bundle of Philodendron with fibers in the center of the vascular bundle and phloem surrounding it. Scalebars: a, b = 100 μm, c, d = 50 μm.
Sclereids may have different forms and sizes (Figure 6a–c). Within the phloem, they are more typically square or polygonal (stone cells) and contain numerous pits and conspicuous pit canals. Holdheid [26] defines that a sclereid is a cell derived from the belated sclerification of a parenchyma cell, and that is in fact the rule in the majority of cases (Figure 6a and b). However, there are lineages in which the sclereids differentiate very close to the cambium (e.g., Pleonotoma, Bignoniaceae, Figure 6c; [20]), and it would be untrue to claim that the derivatives had a stage as a mature parenchyma cell [1]. In these cases, the form is enough to define the sclereid.
Sclereids in the secondary phloem. (a) Sclereids (sc) differentiate from parenchyma cells (arrow) in the nonconducting phloem of Heteropterys intermedia (Malpighiaceae) TS, forming large clusters. (b) Longitudinal radial section of Heteropterys intermedia (Malpighiaceae) showing the sclereid masses. (c) In Pleonotoma tetraquetra (Bignoniaceae), the sclereids differentiate (arrow) close to the cambium within the conducting phloem. c, cambium; pe, periderm; sc, sclereid; sx, secondary xylem. Scalebars: a, b = 400 μm, c = 250 μm.
On the other hand, there are cases where long and slender cells derive from previously mature parenchyma cells and are morphologically difficult to distinguish from fibers. In these cases, these cells are called fiber sclereids and may be even in concentric layers, such as in apple trees and pears (Malus domestica and Pyrus communis, respectively; [15]). Sclereids can also develop with different arrangements in the phloem, being isolated and scattered or in clusters (Figures 6a–c) [5].
The rays in the conducting phloem have typically the same organization in terms of width, height, and cellular composition as the secondary xylem. In this respect the rays vary from uniseriate to multiseriate (Figure 7a) and may be homocellular or heterocellular (Figure 7b). Homocellular rays are those composed of cells of one shape, all procumbent or all upright (common in many shrubs). Heterocellular rays are those where more than one cell shape is present together (Figure 7b). Ray composition is appreciated in radial sections.
Rays in the secondary phloem. (a) Longitudinal tangential section of Brachylaena transvaalensis (Asteraceae) showing storied structure, biseriate to triseriate rays (r), sieve tube elements with simple sieve plate (s) and axial parenchyma cells composed of 4–5 cells, and fibers. (b) Longitudinal radial section of Brachylaena transvaalensis (Asteraceae) showing heterocellular rays (r), with body procumbent and one row of marginal square cells. Fibers (f) in bands. (c) Ray dilatation (rd) by the formation of a dilatation meristem in the center of the ray in Perianthomega vellozoi (Bignoniaceae). f, fiber; p, axial parenchyma cell; r, ray; s, sieve tube element. Scalebars: a, b = 100 μm, c = 300 μm.
Because the vascular cambium produces much more xylem to the inside than phloem to the outside, phloem rays typically greatly dilate toward the periphery of the organ (Figure 7c). It is not uncommon that a dilatation meristem longitudinal to the cambium forms in some barks (Figure 7c), especially in families with very wide, wedge-like rays such as the Malvaceae. Plants with unicellular rays very rarely have dilatation by cell division [15, 26]. Instead, they have great lateral expansion of their single cells. Ray width can be only determined in tangential sections.
Rays are typically exclusively parenchymatic; however, in many species sieve elements appear in the rays and are called ray sieve cells or radial sieve cells [5, 28, 29]. These cells were recorded connecting two different sieve tubes (collections of sieve tube elements). Ray sieve elements seem to be present in taxa where perforated ray cells have been also recorded [30].
The primary phloem derives from the embryo in the seed and the procambium from the organ’s apices. Similarly to the primary xylem, the primary phloem is divided in protophloem and metaphloem (Figure 1d), with the protophloem differentiating first, while the plant is still elongating, and the metaphloem differentiating last. The phloem is always exarch, independently of the organ. Protophloem sieve elements sometimes lack companion cells, such as in Arabidopsis, and in this case the sieve elements are sustained by other neighboring parenchyma cells. Commonly, the protophloem quickly becomes obliterated and loses function. In plants without secondary growth, the metaphloem will be conducted during the entire life of the plant, as in the monocotyledons (Figure 5b–d) [11]. Different vascular plant lineages display different arrangements of the primary xylem and phloem, depending on the stele type. Two main types of steles exist, the protostele and the siphonostele. In the protostele, the entire center of the organ is composed of vascular tissue (Figure 1a), with the phloem in strands alternated with a central xylem in the protostele, haplostele, and actinostele (Figure 1a), while primary phloem is interspersed in the protostele plectostele [6]. The roots of all the vascular plants are protostelic (Figure 1a). The stems, however, can vary. In the lycophytes, they are always protostelic, while in the ferns (monilophytes) they might be protostelic, such as in Psilotum, or in all other range of siphonostelic steles [31]. The siphonostele evolved in concert with the macrophytes and resulted in the formation of a central pith derived from the ground meristem. No lineage displays as much diversity in the primary vasculature architecture as do the ferns. In the seed plants, that is, gymnosperms and angiosperms, the stem stele is always a syphonostele, either a eustele, where discrete vascular bundles form a concentric ring, or the atactostele, a type of stele exclusive of the monocotyledons where the bundles are scattered in the entire stem center. Some lineages of eudicotyledons and Magnoliids have evolved another subtype of siphonostele, the polycyclic eustele, where more than one ring of bundles is present, such as in Piperaceae and Nyctaginaceae.
The primary phloem is simpler than the secondary phloem and is basically formed by sieve elements and parenchyma cells (Figure 1a–d). Fiber caps are commonly present, and they might be phloematic (Figure 5a). For a discussion on their origin, check the section on fibers above. The position of the phloem is typically external or abaxial to the xylem, but in some lineages the bundles are bicollateral (Figure 1b), and phloem is present both inside and outside (abaxial and adaxial), while in amphivasal bundles, the xylem encircles the phloem (Figure 5d), as in the secondary vascular tissues of some Asparagales [32, 33] and Iridaceae corms [34]. In some plant families and orders, intraxylary phloem (perimedullar phloem islands) is a synapomorphy, such as in the order Myrtales and in the families Apocynaceae and Convolvulaceae [35]. These phloem strands are initially primary, but a cambium can differentiate between the protoxylem and the phloem strands and develop secondary tissues inside of the pith.
Being derived from the cambium, the secondary phloem will share a number of characteristics with the secondary xylem. For instance, it is divided in an axial and radial system. The axial system is composed of sieve elements, axial parenchyma cells, and fibers, and the radial system is formed by rays, which are typically parenchymatic (Figure 2a–c). Similar to secondary xylem, the secondary phloem can be storied (Figure 7a) or non-storied (Figure 2b and c), depending whether the cambial mother cells are organized in tiers or not.
Some trees will have growth rings, with an early and a late phloem, both in temperate and tropical regions, but their characterization is only possible with periodical collections [5]. Sometimes, but not always, the fiber band width gives a hint on the presence of growth rings or the formation of very small sieve elements in the late phloem [1, 5].
In conifers (except Gnetales) the secondary phloem is typically marked by an alternation of axial cell types (Figure 3a and b), uniseriate rays, and, in many lineages, axial and radial resin canals (e.g., Pinaceae and Cupressaceae). In the Pinaceae, the phloem is marked by the presence of an alternation of sieve cells and bands of axial parenchyma with phenolic contents, some also with druses. In the nonconducting phloem of Pinaceae, sclereids differentiate. In all other conifers, in addition to the alternation of parenchyma bands and sieve cells, fiber bands are present (Figure 3a and b). Therefore, sieve cells, parenchyma cells with phenolic content, and bands of fibers appear in alternation in non-Pinaceae and Gnetales conifers, including Araucariaceae, Cupressaceae, Podocarpaceae, Taxaceae, and Taxodiaceae [8, 21]. Another marked difference of these conifers compared to Pinaceae is that they contain a lot of crystals in their cell walls, including in Gnetales (see New World Ephedra; [36]), while in Pinaceae they are exclusively inside of idioblastic cells.
In other gymnosperms, in particular in Gnetales and Cycads, the first remarkable difference is the presence of very wide, multiseriate rays alternating with uniseriate rays. The wide rays in both groups have, however, evolved independently, since Cycads are a sister to all other gymnosperms, while Gnetales are within the conifers, as sister to the Pinaceae [31, 37]. In Cyca and the extinct Cycadoidea, sieve cells and phloem parenchyma alternate with fibers, which can be in tangential bands or not [38, 39]. In Cyca, the sieve cells appear in radial rolls [38], while in Cycadoidea there is a constant alternation of one sieve cell or phloem parenchyma to one fiber [39]. The nonconducting phloem of Cycas is marked by the collapse of sieve cells, enlargement of the axial parenchyma cells, ray dilatation, and sclerosis of some parenchymatic cells [38]. More than one ring of secondary phloem is present in some Cycads (e.g., Cycas, Encephalartos, Lepidozamia, and Macrozamia) and Gnetales (e.g., Gnetum), given that they have successive cambia [38, 40].
Within the Gnetales, in Ephedra axial parenchyma cells are interspersed with sieve cells (Figure 4a), and fiber may or may not be present and are typically gelatinous [36]. Fiber sclereids and/or sclereids appear in the nonconducting phloem of other species [13, 22]. In the nonconducting phloem of Ephedra, the sieve cells and Strasburger cells collapse with the enlargement of the axial and radial parenchyma cells (Figure 4a) with more ergastic contents [13]. In Gnetum, large areas of parenchyma sclerify, forming bands in the nonconducting phloem. The secondary phloem of Welwitschia is described as containing a large amount of fibers [21].
Within the angiosperms, the diversity of phloem cell type arrangements reaches its maximum. The structure can be storied (Figure 7a) or non-storied (Figure 2b and c); sclerenchyma can be present or lacking. The rays may be uni-, bi-, or multiseriate. A large array of secretory cells may be encountered, such as resin canals, laticifers, and mucilaginous cells. Crystalliferous parenchyma is also very common, especially when associated with fibers.
The variation in cell type arrangements can be of taxonomic interest. Sieve elements can vary in morphology and arrangement. They can be solitary (Figure 2f), scattered in the phloem (e.g., Eucalyptus, Myrtaceae), in clusters (e.g., Malvaceae; Figures 2a, d and 4c), and in radial or tangential rows (many Bignoniaceae; [20]; Figure 4d). The functional significance of the different arrangements is unknown to date, although this is one of the features in the phloem with the strongest phylogenetic signal.
The presence, type, and arrangements of fibers and sclereids are one of the most informative characters in the bark [4]. In Apocynaceae, the fibers are completely absent, except in Aspidosperma, the sister group of all other Apocynaceae [35]. In Aspidosperma, they can appear solitary scattered across the phloem or in clusters. In some lineages, fibers appear in concentric alternating bands, as in Leguminosae (Papilionoideae), Mimosoideae (Figure 4c) [41], Bignoniaceae [20], and Malvaceae, and this is a constant character among them.
Phloem parenchyma more commonly constitute the background tissue in the phloem but can also be distributed in bands (Figure 4b and c), radial rows, or even only around the sieve tube elements (Figure 4d) [5].
The classic theory of phloem transport is that proposed by Ernst Münch [42], and it involves the formation of an osmotic pressure transport gradient, where certain zones act as sources of sugars (leaves and storage organs), while others act as sinks. Experiments showed that the concentration gradients were always seen to be positive in the direction of flow [43], supporting Münch’s postulate. In a system where transport goes against the direction of transpiration, its functionality relies on the presence of a plasma membrane across the entire system to create an osmotic pressure, hence the need of a conducting system with living cells [44]. Recent studies have been refining aspects involved in the photosynthate conduction to explain long-distance transports across large trees with such a simple system [44, 45]. A direct role of intracellular calcium has also been reported in the dissolution of nondispersive P-proteins and facilitation of transport [46]. Likely, the anatomical structure of the phloem discussed in the previous sections of this chapter will prove to play a role in the system. For instance, phloem sieve element length scale with the tree sizes and sieve plate type [45]. It was also shown that sieve element’s diameter, length, and pore width increase from the top to the base of the trees [47, 48].
Across the entire pathway, sugars are removed from the system to sustain all cells in the plant body. This mechanism is only possible with the concerted mechanism between sieve elements and their close related cells (Strasburger cells and companion cells), with these accompanying cells constantly channeling substances and macromolecules toward the sieve elements [44]. The Strasburger and companion cells carry the loading and unloading of the sieve elements. Given the function of loading and unloading, the companion cell-sieve tube element size ratio is directly related to being in the source or the sink of sugars [44]. For instance, in leaves the companion cells are typically much larger, for they have the high demand of constantly loading the sieve tubes. In areas of release of the sugars (unloading), the companion cells are much smaller or even absent [44].
In the economic uses, it is not always easy to distinguish the use of the phloem from that of the periderm, since both together compose the bark of a woody plant. The phloem corresponds to the inner bark, and the periderm to the outer bark. The bark has a long history of utilization, from the production of remedies [49], aphrodisiacs (yohimbe), insecticides [50], dyes, tannins [50], angostura, fibers [51], gums and resins [50], latex, and flavorings [52].
In indigenous groups from British Columbia (Canada) and Tanzania, barks from dozens of species of woody plants are used as carbohydrate food, medicine, fibers, and structural material [50, 53]. In Mexico the bark of Ficus is used since prehispanic times to create a type of paper called papel amate (from the náhuatl paper = ámatl), used, for example, to create the Aztec codices.
The rubber tree, Hevea brasiliensis (Euphorbiaceae), is known from the extraction of latex to the production of rubber. Laticifers are present in concentric rings in the secondary phloem of the rubber tree and are an important economic asset in some tropical countries. Bark residues have also been considered for mulching [53, 54, 55], to build particle boards [56, 57], as fuel, and a source of food for ruminants [52].
I would like to express gratitude to Ray F. Evert, Veronica Angyalossy, Carmen Marcati, and André C. Lima for allowing their slide collections to be photographed and Leyde N. Nunes for the photo of Tetrapterys leaf, Solange Mazzoni Viveiros for photos of Cucurbita, and Marina Blanco Cattai for picture of Vellozia.
The authors declare no conflict of interest.
Aptamers (Latin word aptus means ‘to fit’ and Greek meros meaning ‘part’) are single stranded oligonucleotides, which act as synthetic ligands for its cognate target molecules. [1, 2] These molecules show high target specificity, selectivity and affinity, which resemble ‘monoclonal antibody’. Similar to antibodies, aptamers also have immense potential to interact with their targets by structural recognition and thus they are termed as ‘chemical antibodies’. [3] Different conformations allow aptamers to bind specifically with their target by ‘lock and key’ model. This hypothesis of binding mechanism is driven by the secondary and tertiary structure of aptamers in-bound state with their targets. Adopting various structures like hairpin loops, bulges, stem-loop, quartets, G-quadruplex, pseudo knots, aptamers can fit into the binding region of the target. [4] Intra and inter molecular interactions like hydrogen bonding, Vander Waals force, hydrophobic interaction, electrostatic forces play major crucial role in aptamer-target interaction. [5] However, the aptamers are primarily synthetic molecules and naturally occurring ribozymes are single stranded RNA molecules, which also have similar recognition domain acting in a similar manner. [6, 7] Aptamers are capable of forming various stable three-dimensional structures in physiological solution. The folding process in solution and the ligand-induced conformational switch is strongly dependent on the presence of divalent cations (magnesium, potassium etc.). [8] There are plethora of computer algorithms enable sequence based modeling of secondary structure of the oligonucleotide aptamers which actually strengthen the predictability of strongest binders with lowest free energy. [9, 10] Aptamers fold into tertiary conformations and bind to their targets through shape complementarity at the aptamer-target interface. [11] An aptamer binds to a protein can modulate protein functions by interfering with protein interaction with natural partners. Similar to antibodies, aptamers can enter to specific target cells via receptor-mediated endocytosis upon binding to cell surface ligands. [12] Importantly, aptamers can penetrate into tumor cores much more efficiently than antibodies due to their ~20–25-fold smaller sizes compared with full sized monoclonal antibodies. [13, 14]
Compared to antibodies, aptamers can be produced using cell-free chemical synthesis and are therefore less expensive for large-scale manufacture. Aptamers exhibit extremely low variability between batches and have better controlled post-production modification, they are minimally immunogenic, and are small in size. (Table 1) The rapidly growing aptamer industry was predicted to reach US $244.93 million by 2020. [15] Presently more than 40 companies are actively engaged in diagnostics and therapeutics research to commercialize these “magic bullets” globally (EU countries, Asia, USA, UK etc.). [16] The largest company is “SomaLogic” (company based on SOMAmer- a patented “Slow Off-rate Modified Aptamer) founded by Prof. Larry Gold at Colorado, USA. Since the advent of aptamers scientists and researchers exploit different applications of aptamers that reflects the following trends in the publications. (Figures 1 and 2).
Monoclonal Antibody | Aptamer |
---|---|
Stability: Monoclonal antibodies require refrigeration to avoid denaturation. Limited shelf life. [17] | Stability: Aptamers do not require refrigeration. Indefinite shelf life. [18] |
Immunogenicity: They can cause immunogenic response. [19] | Immunogenicity: Aptamers are non-immunogenic. [20] |
Production: laborious, expensive, high batch-to-batch variation. | Production: simpler and controlled chemical reactions, little to no variation, automated, chemical synthesis, no contamination. |
Size: Larger in size, they can resist filtration by the kidneys, long half-lives. However, their size prevents access to smaller areas. [21] | Size: Aptamers are small molecules. They are especially subject to kidney filtration, resulting in short half-lives. Compared to antibodies, aptamers can bind to smaller targets. [22] |
Ability to modification: Antibodies cannot accommodate conjugates without negative consequences such as reduced activity. | Ability to modification: Easy to modify, modifications can also be incorporated during synthesis to prevent kidney filtration. [23] |
Comparison between Monoclonal antibody vs. Aptamer:
Publication trend for Search strings: “Aptamers as diagnostics” and “Aptamers as therapeutics” (Source: Scopus).
Publication trend for Search strings: “Aptamers as theranostics” (Source: Scopus).
Back in 1990, two individual groups Prof. Larry Gold and Craig Tuerk from University of Boulder, USA and Prof. Jack Szostak and his student A.D. Ellington from Havard University, USA discovered the evolution process to obtain the oligonucleotide binders and they coined the term ‘Aptamer’ and the process as ‘SELEX’. [24, 25] Systematic Evolution of Ligands by Exponential Enrichment (SELEX) is a common screening process by which aptamers can be selected from an aptamer library which consist of 1024–25 number of various sequences. The method attempts to isolate an aptamer of interest from a pool of randomized library by an iterative cycle of incubation with the target, partitioning and amplification, until the pool of aptamers enriched enough to fit with the target. The SELEX procedure iterates over five basic steps- incubation of aptamer pools with the target, binding, partitioning and washing (to get rid of non-binders which are loosely bound with the target), then elution of positive target-bound aptamers and amplification of enriched pools. Traditionally, the positive pool eluted from last round is being analysed, and high-throughput sequencing is performed.
An array of different RNA and DNA aptamers were isolated against a vast array of targets: ions, [26] low molecular weight metabolites, [27, 28] proteins, [29, 30, 31] sugar moieties [32] lipids, [33] and even whole cells. [34, 35]
To select highly selective, specific aptamers, design of the initial aptamer library is the first and foremost step. In case of determination of the length of the random region researchers should consider the sequence space and structural diversity. The complexity of the initial aptamer library depends on the length of the random window of the aptamer library (If the random window is 40 and if we consider DNA aptamer library, so the complexity of the library is: 4^40 that equals to 1024–25). [36]
Special libraries would consist of specifically designed flanking sequences directing the aptamers to form a specific secondary structure, or include modified nucleotides. In capture SELEX, there is unique docking sequence (12–14 nucleotides long) which enables the library in such a way, that highly sensitive aptamers can be fished out against small molecules. [37, 38] The extended genetic alphabets or combination of artificial xeno nucleic acids (XNA) greatly broaden the diversity of sequences and can influence the properties of the aptamers, such as their in vivo stability or nuclease resistance. [39, 40, 41, 42] Modified nucleotides can be introduced either during the library synthesis or in the post-selection optimization.
In a review article, Maria et al. summarized all key features of designing nucleic acid libraries for SELEX like nature, composition of the library (RNA, DNA or modified nucleotides), the length of a randomized region and the presence of fixed sequences. Different randomization strategies and computer algorithms of library designs were also discussed. [43]
Specific aptamers are screened by the iterative processs of SELEX from a highly diverse pool of oligonucleotides. [44, 45, 46] After the incubation of the random aptamer pool with the target followed by the removal of non- binding aptamers, the bound aptamer species are recovered. These recovered nucleic acid sequences are amplified with PCR (in the case of DNA aptamer) or RT-PCR (for RNA aptamers). In addition to selection against a purified target molecule, SELEX process can be performed against live bacterial cells and even in mammalian cell lines to isolate cancer cell specific aptamers and furthermore it can lead to the identification of novel biomarkers. [47, 48]
A giant advancement of SELEX technology has been made since its discovery in 1990. Conventional SELEX is a well-established and effective method but due to its immense time- and labor-consumption, continuous improvement of alternative methods for aptamer selection has been inevitable.
High throughput SELEX (HT-SELEX), Functional screening (Microfludics or Flow cytometry based SELEX), Cross-over SELEX (where the target is alternatively changing from proteins and cells), (Figure 3) in-vivo SELEX, Spiegelmers selection, de-convolution SELEX are few examples of modern-era screening strategy of aptamers. [49] Cutting-edge functional screening process, the chemical modifications, Next-generation sequencing (NGS technology) enable SELEX more efficient, cost-effective and considerably less time-consuming.
Typical schema of Cross-over SELEX process.
DNA is the backbone of central dogma of our life cycle. Moreover, any form of nucleic acids play a crucial role in our genetic codon. DNA/RNA is an essential bio-macromolecule consist of nucleotide bases such as adenine (A), thymine (T), uracil (U), guanine (G), cytosine (C).
There are various types of modifications (nucleotide base modifications, phosphate backbone modifications, peptide mimic oligonucleotides PNA etc.) available which can prevent aptamers from nuclease degradation. Locked nucleic acid (LNA) is one among them where 2′-oxygen has been linked to the 4′-carbon of the ribose sugar by a methylene bridge, thus completely locking the sugar into a 3′-endo conformation. LNAs increase the thermodynamic stability, binding affinity, and enable the oligonucleotides to prevent serum degradation. [50, 51, 52] These modifications enable the aptamers for biological applications.
Compared to LNAs, the unlocked nucleic acid (UNA) is an acyclic ribose derivative that has increased flexibility. UNAs do not consist the C2’-C3’ bond, which confers the flexibility observed in this modified nucleotide. [53] LNAs increase the melting temperature of the nucleotide by 1–10°C per LNA insertion but UNAs reduce the melting temperature by 5–10°C retaining the nuclear resistance. In case of, Peptide nucleic acid (PNA) in which sugar-phosphate backbone is modified by short stretch of N-(2-aminoethyl)-glycine units connected by peptide bonds, enhances biostability of the modified candidates. [54]
Aptamers have been incorporated in drug development pipeline as they have the capacity to block the downstream signalling (phosphorylation of kinases etc.) of different biomolecules. They can play an important role to regulate various cellular crosstalks. To screen therapeutic aptamers either DNA aptamers or 2′-fluoro modified RNA, a combination of 2′-fluoro pyrimidines and 2′-hydroxyl purines (fYrR) are of major interest. fYrR is the “nuclease stable RNA” and can be easily generated by Y639F modified T7 RNA polymerase. Fovista, an anti-platelet derived growth factor (PDGF) aptamer, was previously DNA aptamer but later modified to augment the stability with the addition of backbone modifications. [55] As with the 2′-fluoro modification, the 2’-OMe modifications adopt a C3’-endo conformation. US FDA approved the first aptamer (Macugen®, pegaptanib sodium) in 2004 against vascular endothelial growth factor for the treatment of age-related macular degeneration. [56] This aptamer was modified with 2′-fluoro-pyrimidines and 2’-O-methyl-purines. The stability of the small aptamer was a critical factor but later which can be circumvented with a 3′-cap and a polyethylene glycol molecule, the half-life of Macugen® was extended to 131 hours at max. [57, 58] Anti-vascular endothelial growth factor (VEGF 165) aptamer Macugen, and an anti-Factor IXa aptamer REG1 were both selected from fYrR libraries, and subsequently 2′-O-methyl nucleosides have been incorporated in order to increase serum stability. [57]
There is a plethora of polymerase enzymes like KOD, Pwo, Phusion, Superscript III, vent (exo-), T7 polymerase have all been shown to be capable of incorporating modified triphosphates into DNA and RNA strands, which open up a new opportunities in aptamer selection strategies. [59] The use of Pfx DNA polymerase allows amplification of Ds-Px base pair in Ex-SELEX protocol where extended genetic alphabets were included in complexity of nucleic acid library. [60]
Several limitations of aptamers should be considered in the process of in-vivo applications of nucleic acid aptamers. Being polynucleotides, nucleic acid aptamers are naturally susceptible to enzymes degradation by exo and/or endo-nucleases, leading to a reduced in vivo circulatory half-life. This drawback can be alleviated by side chain chemical modifications to aptamers, incorporating unnatural nucleotide bases (locked and unlocked nucleic acids) and capping the aptamer ends, thus minimizing the susceptibility to endonuclease and exonuclease attack. [50, 51] Short blood residence time is another challenge with in vivo aptamer applications, which is due to fast removal of aptamer from the circulation by renal filtration as most aptamers have a size smaller than the renal filtration threshold of 40 kDa. [31] To achieve desired serum half-life, aptamers can be engineered by conjugation with a terminal polyethylene glycol (PEG), although this may compromise the extent of tumor penetration [61]. In some cases, post-SELEX modifications following the selection of aptamers may alter the 3-D structure of the aptamers, leading to the lost or altered binding affinity and specificity. Such problems can be prevented by using random aptamer pools containing modified nucleotides during the SELEX process. [62, 63] In addition, the ability of aptamers to interact with cells may decrease due to repulsion of nucleic acids by negatively-charged cell membranes. This can be refuted by increasing the binding affinity and specificity of aptamers toward their cell surface receptors to trigger receptor-mediated endocytosis.
In the field of oncology, two aptamers, namely, AS1411 and NOX-A12, have entered clinical trials. [45, 64] AS1411 (formerly ARGO100; Antisoma) is a guanine quadruplex aptamer obtained from a guanine-rich oligonucleotide library in the anti-proliferation screen, which is not a typical SELEX process. [65] The guanine quadruplex structure benefits AS1411 because it is resistant to nuclease degradation and enhances cell uptake. In in-vitro validations, AS1411 inhibits more than 80 types of cancer cell lines. In addition, the target of AS1411 has been verified to be nucleolin, and the relevant mechanism and specificity against cancer cells have also been described. In preclinical tests, AS1411 shows efficacy toward xenograft models, including non-small cell lung, renal, and breast cancers. It entered clinical trials in 2003 and passed phase II trials for acute myeloid leukemia. A subsequent phase II trial for renal cell carcinoma started in 2008 (clinical trial ID NCT00740441). [66] NOXA12 (Olaptesed pegol; Noxxon) is an L-form RNA aptamer known as Spiegelmer and is used for cancer therapy. NOX-A12 can bind to its target chemokine CXCL-12 and blocks its interaction with its receptor. [67] This disrupts the tissue gradient of CXCL-12 and reduces the cancer cell homing that might lead to tumor metastasis and drug resistance. [68] Currently, phase II clinical trials for NOX-A12 are underway for the treatment of chronic lymphocytic leukemia and refractory multiple myeloma (clinical trial IDs NCT01486797 and NCT01521533). [67] Aptamer based cancer therapeutics have immense potential for precise and less toxic treatment for cancer patients. [46]
Aptamers can be used in-vitro and in-vivo as well. [69] In terms of in vivo diagnostics, ‘escort’ aptamers can be implied as vehicles for a detectable molecules, such as radionuclides, fluorophores etc. [70, 71, 72] The development of new agents like radio-pharmaceuticals is challenging. There are some important factors such as efficiency of the radiolabeling process, specific activity (radioactivity per moles e.g. Ci/μmol), chemical purity, radiochemical and chemical stability and shelf life of the final product. [73] Mostly, radiolabeling strategies for aptamers are similar as for proteins, or antibodies. Aptamers can be easily chemically modified at its 5′ or 3′ end with a desired functional group for radiolabeling (Figure 4).
Aptamers modified with radiolabelled molecules for disase diagnosis (Figure adapted from Gijs et al) [73].
Radiohalogens (fluorine-18, bromine-76, iodine-125 etc.) are the most commonly used for radiolabelling oloigonuclotides which are often accompanied with prosthetic groups. Recently, click-chemistry for radiofluorination was demonstrated on antisense oligonucleotides and siRNAs. [74, 75] Another report used photoconjugation as strategy for the radiofluorination of an aptamer. [76] Oligonucleotides have also been radiolabeled with the radiohalogens such as bromine-76 for PET imaging and iodine-123 for SPECT (Single photon emission computed tomography) imaging. In addition, iodine-125 has been used to radiolabel antisense oligonucleotides, aptamers and spiegelmers for theranostic applications. Due to the harsh and non-aqueous reaction conditions usually needed to radiolabel prosthetic groups, it is performed before the conjugation process to the oligonucleotide. [73]
Till date, a plethora of aptamers have been modified or labelled with radioactive molecules. Aptamers against several important biomarkers like PMSA, Tenascin C, thrombin, MUC1 were already exploited for radiolabelling. Aptamer-based radiopharmaceuticals were primarily developed for imaging and therapy of cancer diseases, metabolic disorders and others. The aptamers are mainly radiolabeled with technetium-99 m for SPECT (Single photon emission computed tomography), PET (Positron emission tomography) imaging. Very few aptamers were published related to PET imaging, and there is only one study of radiolabeled aptamers for therapy by Bandekar et al. [77] Other radiolabeled aptamers have only been tested for preclinical applications or in the course of preclinical assesement.
Molecular nuclear imaging technique is a diagnostic process of non-invasive visualization of any disease in-vivo at molecular level with high precision. For nuclear imaging, the probes used for radiolabelling has to be modified accordingly. Aptamers are the most promising candidates with versatile modification capibility, can be easily engineered for various imaging and other diagnostic purposes.
The first radiolabeled aptamer for nuclear imaging was discovered by Charlton et al. A DNA aptamer, NX21909, was selected against human neutrophil elastase, an enzyme which is secreted by neutrophils and macrophages during inflammation to kill pathogens. [78]
Aptamer TTA1, an RNA aptamer targeting the extracellular matrix protein tenascin C (TN-C), was the first radiolabeled aptamer which was used as molecular cancer imaging agent. Aptamer TTA1 was generated by a cross-over SELEX involving the purified recombinant TN-C protein and TN-C-positive U251 glioblastoma cells. [79, 80]
There are a unique group of aptamers (generally RNA aptamers) which can bind specifically with their cognate fluorogen molecules like DFHBI, thiazole orange, thioflavin T etc. [49, 81, 82] The non-fluorescent moelcules (native unbound state) become fluorescent (bound state) after binding to the aptamers and these “light-up” aptamers generate fluorescence signal. In the omni-presence of target molecules (small pre-miRNAs) and malachite green (fluorogen) light up aptasensors ‘malaswitch’ exhibit fluorescence enhancement. [83] We can engineer the small-molecule specific aptamers (like aptamers for some pesticides, toxins, small metaboltes) in such a way, that combined with light-up aptamers, they can generate a detectable signal. Light up aptasensors are promising alternative biosensor for label free sensitive detection of small molecules. [84]
With more focus on in vivo studies for potential clinical applications, aptamers can be developed in combination with DNA nanostructures, nanomaterial, or microfluidic devices as diagnostic probes or therapeutic agents for cancers, infectious diseases, genetic, metabolic, neurological disorders, lifestyle diseases and several others. The use of aptamers as targeting agents in drug delivery can also be explored. Aptamers might be exploited to develop portable, low-cost and robust diagnostic kit using simple devices for real-time and on-site POC (point-of-care) detection and monitoring, instead of the laborious and time-consuming diagnostic tests currently available only in clinical labs. Regarding therapeutics approach, there is still untapped potential in the combination of the target recognition and strong binding property of aptamers with exquisitely designed nanomaterials. It can be used as an effective alternative drug delivery platform. Variety of materials such as liposomes, polymer vesicles and silica nanoparticles, combined with DNA/RNA aptamers, has shown feasibility for use in in vivo targeted drug delivery. [85, 86] The integration of diagnostic capability with therapeutic interventions termed, as “Theranostics” is critical to address the challenges of disease heterogeneity and adaptation. Although aptamers have immense potential as theranostic agents, tailor-made modifications, validation of experiments need to be executed before aptamer-based drug delivery can reach clinical trials and eventually the patient management system.
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